**Screening of Bacterial Quorum Sensing Inhibitors in a** *Vibrio fischeri* **LuxR-Based Synthetic Fluorescent** *E. coli* **Biosensor**

**Xiaofei Qin 1,2, Celina Vila-Sanjurjo 2,3, Ratna Singh <sup>4</sup> , Bodo Philipp <sup>5</sup> and Francisco M. Goycoolea 2,6,\***


Received: 13 August 2020; Accepted: 18 September 2020; Published: 22 September 2020

**Abstract:** A library of 23 pure compounds of varying structural and chemical characteristics was screened for their quorum sensing (QS) inhibition activity using a synthetic fluorescent *Escherichia coli* biosensor that incorporates a modified version of lux regulon of *Vibrio fischeri*. Four such compounds exhibited QS inhibition activity without compromising bacterial growth, namely, phenazine carboxylic acid (PCA), 2-heptyl-3-hydroxy-4-quinolone (PQS), 1*H*-2-methyl-4-quinolone (MOQ) and genipin. When applied at 50 µM, these compounds reduced the QS response of the biosensor to 33.7% ± 2.6%, 43.1% ± 2.7%, 62.2% ± 6.3% and 43.3% ± 1.2%, respectively. A series of compounds only showed activity when tested at higher concentrations. This was the case of caffeine, which, when applied at 1 mM, reduced the QS to 47% ± 4.2%. In turn, capsaicin, caffeic acid phenethyl ester (CAPE), furanone and polygodial exhibited antibacterial activity when applied at 1mM, and reduced the bacterial growth by 12.8% ± 10.1%, 24.4% ± 7.0%, 91.4% ± 7.4% and 97.5% ± 3.8%, respectively. Similarly, we confirmed that *trans*-cinnamaldehyde and vanillin, when tested at 1 mM, reduced the QS response to 68.3% ± 4.9% and 27.1% ± 7.4%, respectively, though at the expense of concomitantly reducing cell growth by 18.6% ± 2.5% and 16% ± 2.2%, respectively. Two QS natural compounds of *Pseudomonas aeruginosa*, namely PQS and PCA, and the related, synthetic compounds MOQ, 1H-3-hydroxyl-4-quinolone (HOQ) and 1H-2-methyl-3-hydroxyl-4-quinolone (MHOQ) were used in molecular docking studies with the binding domain of the QS receptor TraR as a target. We offer here a general interpretation of structure-function relationships in this class of compounds that underpins their potential application as alternatives to antibiotics in controlling bacterial virulence.

**Keywords:** compounds screening; quorum sensing inhibition; antibacterial; molecular docking

#### **1. Introduction**

Bacteria communicate with secreted chemical signalling molecules that act as autoinducers in a phenomenon known as quorum sensing (QS) which allows them to fulfil a variety of functions, including bioluminescence, virulence and biofilm formation, among others [1]. In the case of Gram-negative bacteria, the autoinducer molecules belong mostly to the family of the acyl homoserine lactones (AHLs), which accumulate to particular threshold concentrations once the population of cells grow to sufficient [2,3].

AHL synthesis relies on the synthase LuxI family, while AHL reception depends on LuxR-type transcriptional regulators, which include the nominal LuxR protein from *Vibrio fischeri*, but also the related TraR and LasR from *Agrobacterium tumefaciens* and *P. aeruginosa*, among others. These act as transcriptional activators or repressors of the target QS genes [4–7]. The canonical LuxR protein from *V. fischeri* comprises two domains. The N-terminal domain is responsible of AHL binding and also can mediate protein dimerisation [8]. In contrast, the C-terminal domain contains a helix-turn-helix (HTH) motif, which is thought to make sequence-specific binding to DNA and to drive RNA polymerase binding to target promoters [9]. Much research has been conducted in recent years on mutant- LuxR-type proteins [10–16]. Among these, the TraR protein from *Agrobacterium tumefaciens* has been thoroughly studied, and its crystal structure has been solved, revealing the presence of the above mentioned two functional domains. In TraR, the N-terminal domain binds to N-3-oxooctanoyl-L-homoserine lactone (OOHL), and the C-terminal domain interacts with the DNA binding domain of the tra box [7,17]. TraR is a dimer in the presence of OOHL, and the TraR-OOHL-*tra* DNA ternary complex can be used as a prototype for the large family of AHL-induced transcription activators. The LasR protein of *Pseudomonas aeruginosa* shares 70% homology with TraR of *A. tumefaciens*, and the 3D model of the TraR active site closely resembles the X-ray structure of the LasR active site [18]. The signal binding sites in both apo-proteins are highly accessible, so TraR constitutes a useful model receptor which allows predicting the ability of putative QS inhibitors (QSIs) to block QS-based mechanisms in the human pathogen *P. aeruginosa* [19,20].

The development of strategies aimed to block or disrupt QS is gaining momentum. These efforts are directed to inhibit the production of virulence factors at the site of infection by dismantling the collective power of bacterial pathogens, an approach known as quorum quenching (QQ) or QS inhibition [21,22]. In the last years, QSIs have attracted significant attention from the scientific community and are considered as potential weapons and new generation antimicrobials in the therapeutic arsenal against infections caused by drug-resistant bacteria [23–25].

There are three major approaches to target bacterial QS using QSIs: (i) destruction of the signalling molecule, (ii) inhibition of signal production, and (iii) inhibition of the receptor [26]. To the date, most of the literature on QQ is centred in the investigation of AHL-degrading enzymes [27] and more abundantly, in QSIs that targeted to specific QS regulators of diverse species [26]. These QSIs may structurally resemble the natural ligand or may on the contrary, have an entirely different molecular structure [28,29]. QSIs include compounds of diverse sources, both natural and synthetic, and include fungal metabolites, plant substances, antibiotics, and synthetic derivatives of QS autoinducers or natural antagonists, to name a few [26–29]. Identification of QSIs is commonly performed through the screening of compound libraries and biosensor-based analysis of the QS response alone or combined with computer modelling analysis. These methods have allowed expanding the catalogue of available QSIs, which includes a variety of AHL analogues, brominated furanones, polyphenolic compounds and polypeptides [23,26,30–32].

Regarding computer modelling, docking-based screening of candidate QSIs is normally carried out by using a genetic algorithm on a library of 2344 compounds and calculates their binding free energy, hence identifying those candidates able to interact with target conserved residues in the binding site of a LuxR-type model receptor [33]. In-silico screening of ligand databases has thus become an important strategy towards the discovery of novel QSIs. GRID molecular interaction fields (GRID-MIFs) is accepted as an efficient method in virtual screening of candidate molecules which target protein binding sites. It is a computational procedure for detecting energetically favourable binding regions in proteins and small molecule drugs of known 3D structure. The energies are calculated using the electrostatic, hydrogen-bond, Lennard Jones, and entropic interactions of chemically selective probes with the chosen biological target. The program works by defining a three-dimensional grid of points that contains the chosen substrate binding site. The above mentioned calculations are repeated for each node in the three-dimensional grid and for each probe being considered. The results of these calculations are a collection of three-dimensional matrices, one for each probe-target interaction. A detailed description of the GRID program, the force field parameters, and details of calculations can be found elsewhere [34,35]. Briefly, a grid is projected inside the protein regions and cavities of interest. The probes are functional groups that can move stepwise from grid point to grid point. The calculated interaction energies of the probes are computed to create the MIFs, which represents the potential interaction of the protein with a particular chemical group. GRID-MIFs is considered as a high-throughput screening method to virtually analyse protein-ligand interactions [36,37].

In this study, we have screened 23 potential QS inhibition compounds, representing five groups according to their chemical structures. We found that seven of them were potent, dose-dependent inhibitors of the *V. fisheri* LuxR-based system expressed on recombinant *E. coli* biosensor cells, including two unprecedented ones. Another five compounds have shown antibacterial activity, while the remaining eleven compounds were inert at the tested doses. Moreover, we performed in silico GRID-MIFs-based computational molecular docking on the 3D crystal structure of TraR that allowed us to propose a hypothesis on the disparate effects observed experimentally for compounds of chemically related structure. We propose that our biosensor- and docking-based approaches have a high potential for drug designing purposes.

#### **2. Results**

#### *2.1. Screening a Panel of Potential Quorum Sensing Inhibitors*

We selected a panel of 23 pure compounds with the potential to act as inhibitors of LuxR-AHLmediated QS. These comprised the following groups, namely: Group (1) lactone analogues, Group (2) aromatic ring structures, Group (3) heterocyclic compounds, Group (4) *Pseudomonas spp-* relevant compounds, and Group (5) structurally unrelated compounds. Figure 1 shows the chemical structures of the cognate AHL molecules of LuxR and TraR regulators, namely *N*-3-oxohexanoyl-L-homoserine lactone (3OC6HSL) and OOHL, respectively.

All 23 compounds were tested using the *E. coli* Top10 pSB1A3-BBaT9002 biosensor, which expresses a synthetic version of the *lux* regulon of *V. fischeri* and produces a fluorescent as a response to external (3OC6HSL) [38]. The rate of the density-normalised fluorescence of the *E. coli* biosensor as a function of the 3OC6HSL concentration displays a Hill behaviour, with a *k*Hill of 7.48 × 10−<sup>10</sup> ± 9.03 × 10−<sup>11</sup> M. At 3OC6HSL concentrations higher than 1 × 10−<sup>9</sup> M, the fluorescence response is saturated, while the fluorescence levels are undetectable at a 3OC6HSL concentration of 1 × 10−<sup>10</sup> M [38]. We evaluated the QS inhibitory effect of our panel of compounds by their ability to reduce the fluorescence response of this *E. coli* biosensor without compromising cell growth.

**Figure 1.** *Cont*.

**Figure 1.** Chemical structures of studied compounds. Group (1) lactone analogues, Group (2) aromatic ring structures, Group (3) heterocyclic compounds, Group (4) *Pseudomonas* spp.-relevant compounds, and Group (5) structurally unrelated compounds. In the Figure are also shown the structure of natural LuxR and TraR ligands, namely 3OC6HSL and OOHL. Other details of the series of compounds are given in Materials and Methods section.

Screening of the 23 compounds revealed that they could be classified into three main categories based on their QS inhibition activity, relative to their effects on bacterial growth. The first category refers to compounds that have not shown QS inhibition nor antibacterial activities (i.e., no apparent effect on the fluorescent response and growth of the *E. coli* biosensor). The second includes compounds with the ability to reduce the QS response of the biosensor without compromising cell growth. While the third one comprises compounds with the ability to reduce the QS response at the expense of hampering cell growth. Figure 2 shows representative dose-response curves of compounds belonging to the classes 1, 2, 3, namely gardenoside (13) (Figure 2a,d,g), caffeine (11) (Figure 2b,e,h) and furanone (4) (Figure 2c,f,i) on the fluorescence (Figure 2a–c), growth (Figure 2d–f) and density-normalized fluorescence (Figure 2g–i) of the *E. coli* biosensor. A selection of dose-response effects of other compounds is available as Supporting Information, as discussed below.

**Figure 2.** Effect of increasing concentrations of gardenoside, caffeine and furanone on the fluorescence (**a**–**c**), growth (**d**–**f**) and density-normalised fluorescence (**g**–**i**) of the *E. coli* biosensor over time.

The three compounds are chosen as representatives of the following categories: no inhibition (gardenoside; **a**,**d**,**g**), QS inhibition in the absence of growth reduction (caffeine; **b**,**e**,**h**) and QS and growth inhibition (furanone; **c**,**f**,**i**). Results from additional experiments on other compounds are available in Supporting Information. Data shows the mean and standard deviation of a representative experiment with triplicated treatments.

Next, we calculated the end-point effect of the 23 candidate compounds on the QS-based response and growth of the biosensor, relative to those of untreated cells. Figure 3 shows end-point, relative reduction of density-normalised fluorescence and cell density of compounds tested at fixed concentrations of 5 × 10−<sup>5</sup> M (Figure 3a,b) and 1 × 10−<sup>3</sup> M (Figure 3c,d). The rationale behind the chosen concentrations was based on the maximum water solubility of the compounds. Relative values close to 1.0 indicate none or negligible effect of a given compound on the density-normalised response and/or growth of the biosensor. In turn, we considered that relative fluorescence values significantly lower than 1.0 as were a diagnose of inhibition of the QS-based response, proven that the relative OD<sup>600</sup> values stayed close to 1.0. This condition applied to vanillin (**5**), caffeine (**11**) and *trans*-cinnamaldehyde (**6**) applied at 1 × 10−<sup>3</sup> M (Figure 3a,b); and to phenazine carboxylic acid (PCA, **19**), 2-heptyl-3-hydroxy-4-quinolone (PQS, **14**), genipin (**12**) and 1*H*-2-methyl-4-quinolone (MOQ, **15**) applied at 5 × 10−<sup>5</sup> M. In all cases, the relative QS and OD<sup>600</sup> data were compared statistically against itaconic acid, due to its negligible effect on both parameters. − − − −

− **Figure 3.** End-point effect of the 23 candidate compounds on the QS-based response and growth of the *E. coli* biosensor. (**a**) Effect of compounds **10, 2, 9, 3, 1, 7, 8, 13, 21, 5, 11, 6, 4, 22**; applied at 1 × 10−<sup>3</sup> M on the density-normalised fluorescence of treated cells relative to control cells.

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Relative fluorescence was calculated as follows: the mean of the last ten values of density-normalised fluorescence over time, corresponding to 246–300 min of incubation (see Figure 2) was divided by the corresponding values of untreated cells. (**b**) Effect of compounds **10, 2, 9, 3, 1, 7, 8, 13, 21, 5, 11, 6, 4, 22;** applied at 1 × 10−<sup>3</sup> M on cell density of treated cells relative to control cells. Relative OD<sup>600</sup> was calculated as follows: the mean of the last 10 OD<sup>600</sup> values over time, corresponding to 246–300 min of incubation (see Figure 2) was divided by the corresponding values of untreated cells. (**c**)**.** Effect of compounds **21, 17, 16, 23, 19, 14, 12, 15, 18, 20;** applied at 5 × 10−<sup>5</sup> M on the density-normalised fluorescence of treated cells relative to control cells. Relative fluorescence was calculated as in (**a**,**d**) Effect of compounds **21, 17, 16, 23, 19, 14, 12, 15, 18, 20**; applied at 5 × 10−<sup>5</sup> M on cell density of treated cells relative to control cells. Relative OD<sup>600</sup> was calculated as in (**b**) *t*-Student statistical comparisons were made using itaconic acid as a reference treatment (\* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001 and \*\*\*\* *p* < 0.0001). Data show the mean and standard deviation of three independent experiments with triplicated treatments.

Figure 3a shows that the compounds *trans*-cinnamaldehyde (**6**) (*p*<0.0001), caffeine (**10**) (*p*<0.0001), and, to a less extent, vanillin (**5**) (*p* < 0.001) can significantly reduce the QS-based fluorescence of the biosensor when applied at 1 × 10−<sup>3</sup> M. Importantly, the QS inhibitory effect of caffeine (10) was not accompanied by any compromise of cell growth (Figure 3b). As for vanillin (**5**) (*p* < 0.01) and *trans*-cinnamaldehyde (**6**) (*p* < 0.01), they slightly reduced cell growth by 16.0% and 18.6%, respectively (Figure 3b). Furanone (**4**) (*p* < 0.0001), and polygodial (21) (*p* < 0.0001), which abolished the QS-based fluorescence of the biosensor at the tested concentration, they concomitantly exerted a dramatic antibacterial effect (cf. Figure 3a,b). Figure 3c shows that compounds PCA (**19**) (*p* < 0.01), PQS (**14**) (*p* < 0.001), genipin (**12**) (*p* < 0.001) and MOQ (**15**) (*p* < 0.0001) significantly reduced the density-normalized fluorescence of the biosensor when tested at 5 × 10−<sup>5</sup> M (Figure 3a) and, importantly, they did not exert a significant effect on bacterial growth (Figure 3b). By contrast, compounds PYO (**18**) (*p* < 0.0001) and PMS (**20**) (*p* < 0.0001), showed a strong deleterious effect both on the QS response of the biosensor and its growth (cf. Figure 3c,d). We carried out a series of experiments to confirm that inhibition of the density-normalised fluorescence observed with PYO (**18**) and genipin (**12**) at sub-lethal doses was in fact due to interference with the QS response of the biosensor and not due to GFP fluorescence quenching. To this end, we used a control *E. coli* strain Top10 pBCA9445-jtk28282::sfGFP, which constitutively expresses a super folded version of GFP (sfGFP). We compared the effects of both compounds in the *E. coli* biosensor and control strains (Figures S2–S8). In our hands, neither of them at concentrations in the range 1 × 10−<sup>8</sup> M to 1 × 10−<sup>4</sup> M (Figures S3–S5) acted as quenchers of sfGFP fluorescence in the control strain *E. coli* Top10 pBCA9445-jtk28282::sfGFP. We observed a high degree of experimental variability and not apparent dose-response effects of PYO at sub-lethal concentrations ranging from 1 × 10−<sup>8</sup> to 1 × 10−<sup>4</sup> M (Figure S5). The implications of these disparate observations are further considered in the Discussion section.

#### *2.2. Computation of GRID-MIFs*

The three-dimensional molecular crystal structure of TraR receptor was obtained from the protein data bank (PDB No. 1L3L). Information about the binding region where the natural ligand OOHL interacts with TraR was described elsewhere [17]. Favorable interaction points at the binding site of the receptor was studied with GRID-MIFs. To define the GRID maps at TraR binding site, the Autogrid utility inbuilt in AutoDockTools 1.5.6 software was applied. Three different chemical probes (HD, HA, DRY) were used, representing the potentially significant functional group at the binding site. In Figure 4a the GRID-MIFs generated at TarR binding site is shown. Here, the green patch generated by DRY probe accounts for the favourable hydrophobic interaction ligand and receptor; blue contours were generated by HD (donor) probe responsible for favourable hydrogen bonds between receptor and ligand, and red contours were generated by HA (acceptor) probe, which informs about favourable hydrogen bonds between ligand and receptor amino acid residues. Comparing with the natural substrate OOHL (Figure 4b) the dry probe matches with the ring and all carbons in the substrate, the HD (blue) probe matches the NH group of substrate and HA (red) matches the O functional group present in the substrate.

**Figure 4.** (**a**) GRID-MIFs for TraR protein with DRY probe (green) showing favorable hydrophobic interaction sites. GRID-MIFs for TraR protein with HD (blue) and HA (red) probes showing favorable hydrogen bond (blue for hydrogen bond donor and red for hydrogen bond acceptor) binding sites. (**b**) Natural substrate OOHL.

#### *2.3. Docking and Interaction of Selected Compounds*

We docked six selected compounds, based on their generated GRID-MIFs, over the binding domain of TraR. We included the cognate OOHL ligand as a reference. This enabled to gain Information on dock score, hydrogen bond and hydrophobic interactions. We selected ligand conformations using that of the natural ligand OOHL into the binding site as a reference (Figure 5a). Thus, we fixed docked ligand binding poses to that of the natural ligand. In other words, superimposition of the docked ligand on OOHL conformation in TraR's crystal structure implied that the aromatic ring of the ligand coincided with the lactone ring of the natural substrate. Table 1 lists the estimated binding energy values for the docked compounds. Strikingly, PCA (19) and PQS (14) showed a more negative docking score (i.e., predicted free energy of binding) than that estimated for the cognate ligand OOHL, which would in principle indicate a stronger affinity of these two compounds towards the binding site of TraR (Table 1). Figure 5b shows the docking-predicted binding pose of PCA into TraR's binding pocket. Two ligand conformations were identified for PQS (14), namely PQS-conf A (Figure 5c) and PQS-conf B (Figure 5d and Table 1). Both conformations were similar to OOHL binding pose, only differing in ring flip, which affects hydrogen bond interactions between ligand and receptor (cf. Figure 5a–c). The estimated docking score for PQS (14)-conf A was more negative than that of OOHL, whereas the predicted value for PQS-conf B was less negative, indicating a potential stronger affinity of conf B vs conf A towards the binding site of TraR (Table 1). Docking scores of smaller ligands, namely MOQ, HOQ and MHOQ were less negative than that of OOHL, predicting poorer affinities for these ligands towards TraR (Table 1).

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**Figure 5.** Interaction of TraR receptor with (**a**) OOHL; (**b**) PCA, (**c**) and (**d**) PQS; (**e**) MOQ; (**f**) HOQ and (**g**) MHOQ. (**c**) shows us the PQS conf A and (**d**) shows PQS conf B. Close-up view of all compounds binding site, oxygen carbon and nitrogen are colored in red yellow and blue, respectively, hydrogen bonds are shown as blue line.


**Table 1.** Cumulative results for the compounds docked against TraR protein.

γ γ γ γ α γ γ We studied in detail the docking-predicted poses of selected candidates to elucidate the mechanisms of receptor-ligand possible interactions (Table 1). Figure 5 shows interaction maps of the six selected compounds (Table 1) with TraR's binding pocket. Clear from Figure 5a are the presence of four hydrogen bonds established between the cognate substrate OOHL and specific amino acids at the binding pocket of the receptors The H- bonds are located as follows: (1) between Trp 57 in TraR and keto group present in the lactone ring, (2) between Tyr 53 in TraR and the 1-keto group of the lactone, (3) between Asp 70 in TraR and the imino group in OOHL and (4) between Tyr 61 in TraR and the 3-keto group of OOHL (Figure 5a). Apart from these bonds, the cognate ligand also establishes strong

γ

hydrophobic interactions with nearby residues, as shown in Table 1. Altogether, it seems that both hydrophobic and hydrogen bond interactions are important for ligand selectivity. Our docking results confirm previous docking predictions of binding and pose conformations for OOHL in TraR's binding site [18]. Figure 5b shows the docking-predicted map for PCA (19)-TraR interaction (19). Importantly, our predictions show the presence of three hydrogen bonds with TraR's binding pocket, namely: (1) between Trp 57 in TraR and the deprotonated N present in the aromatic group of PCA, (2) and (3) between Thr 129 in TraR, an intermediate water molecule and the carboxylic acid functional group of PCA (Figure 5b). Again, our docking predicted several hydrophobic interactions between specific aminoacids of the binding site of TraR and the aromatic and heterocyclic rings of PCA (19) (Table 1). Figure 5c shows the predicted maps for PQS-confA and the binding site of TraR. In this case, two hydrogen bonds are predicted to form, namely: (1) between Thr129 and the keto group present in the aromatic ring of PQS (14), (2) between a hydroxyl group in PQS and a molecule of water (Figure 5c). In turn the predicted map for PQS-conf B and TraR interaction (Figure 5d shows one hydrogen bond formed between Trp 57 in TraR and the keto group present in the aromatic ring ofPQS (14). Both PQS-conf A and -confB are expected to establish strong hydrophobic interactions with the binding pocket of TraR, as listed in Table 1 and the long carbon chain of PQS in both poses cover most of the hydrophobic patch generated by GRID (cf. Figures 4 and 5c,d). Panels e, f and g of Figure 5, illustrate the prediction interaction maps for TraR and MOQ (15), HOQ (16) and MHOQ (17), respectively. Docking-predicted poses in the binding site of the receptor are very similar (cf. Figure 5e–g); hence, the predicted interaction sites for the three of them did not very much. However, in the case of both MOQ (15) and MOQH (17) an hydrogen bond was predicted between Trp 57 in TraR and the keto group in the aromatic ring of the ligands (cf. Figure 5e,g), while this interaction was not observed in HOQ (15) (Figure 5f). Strikingly, a relative inhibition of the QS-based response of the *E. coli* biosensor was found to be significantly higher for MOQ (15) than that of HOQ (16) and MOQH (17) (Figure S9). This, in principle would indicate that the specific position of H-bonding between these kind of molecules and the binding site of TraR would not necessarily explain *per se* towards LuxR-based proteins.

#### **3. Discussion**

QSIs are gaining momentum as potent alternatives to the use of classical antibiotics in the context of rising resistance spread and with enormous potential in many fields, from food science to agriculture and medicine [39,40]. QSIs are structurally diverse molecules that can be synthesised or extracted from natural sources. Here, we screened a total of 23 chemically diverse compounds with potential QS inhibition activity to identify their impact on LuxR-regulated QS models. To this end, we used a well-established *E. coli* Top10 biosensor assay and in silico modelling to recreate the structural interactions between the ligand candidates and the LuxR-like receptor.

The candidate compounds were classified into five broad groups according to with their chemical structure (Table 1). Group 1 comprised the alkyl-substituted lactones (Table 1): γ-valerolactone (**1**), L-homoserine lactone (**2**), α-methyl-γ-butyrolactone (**3**), and furanone **4**. In this group, only furanone (4) showed a significant inhibition (≥37.5%) of the fluorescent- QS-based response of the biosensor at concentrations ≥ 5 × 10−<sup>6</sup> M 37.5% (Figure 3a and Figure S10). At 1 × 10−<sup>3</sup> M, furanone completely abolished the QS-based response of the biosensor, albeit a dramatic, deleterious effect on cell growth (Figure 3b and Figure S10).

γ-Butyrolactones were the first class of small signalling molecules identified from Gram-positive species *Streptomyces griseus*. They were found to induce streptomycin production and sporulation [41]. Even though the chemical structure of γ-butyrolactones is rather similar to that of AHLs, except for the carbon side-chain, it is known that the receptors of γ-butyrolactone do not bind to AHL receptors and that AHL do not bind to γ-butyrolactone receptors. Also, both receptors have low structural similarity [41]. The functions of the signalling molecules also differ as AHLs i.a. are known to regulate QS in Gram-negative bacteria, while γ-butyrolactones mainly regulate the production of antibiotics and differentiation.

From our results, we found that neither α-methyl-γ-butyrolactone, γ-valerolactone nor L-homo-serine lactone showed any QS inhibition activity. These results are consistent with the notion that the carbon side chain is important for binding to the LuxR receptor. Also, it is not unexpected that γ-valerolactone did not show activity, as the methyl substituent is in a different position of the lactone ring than that in 3-oxohexanoylhomoserine. As suggested in previous studies, our results support the proposal that the LuxR receptor does not recognise α-methyl-γ-butyrolactone nor other related lactones (e.g., γ-valerolactone).

Halogenated furanones which resemble AHLs structurally were first discovered in the marine red alga *Delisea pulchra* and proved to preventing swarming motility with the opportunistic human pathogen *Proteus mirabilis* and *S. liquefaciens* without affect growth rate [42,43]. These compounds were the first QSIs found to occur in nature. After that, Manefield et al. further obtained proof-of-principle of the QS inhibition activity of the same type of halogenated furanones by specifically interfering with AHL-mediated gene expression at the level of the LuxR protein [44]. Furthermore, Ren et al. found that (5*Z*)-4-bromo-5-(bromomethylene)-3-butyl-2(5*H*)-furanone is a non-specific intercellular signal antagonist [45]. Later, Defoirdt et al. proved that this furanone disrupts quorum sensing-regulated gene expression in *Vibrio harveyi* by decreasing the DNA-binding activity of the transcriptional regulator protein LuxR [46]. Besides halogenated furanones with side alkyl chains that have QS inhibition activity, some synthetic halogenated furanones lacking alkyl chain also have anti-biofilm formation activity through interference with QS, and have been found to protect surfaces from being colonised by *S. epidermidis* [47]. Compound (5*Z*)-4-bromo-5-(bromomethylene)-3-butyl-2(5*H*)-furanone included in our library, a synthetic derivate of natural furanone has been reported as a potent antagonist of bacterial QS that not only can increase bacterial susceptibility to tobramycin and SDS when it is applied for action with *P. aeruginosa* biofilms but also can inhibit virulence factor expression because of targeting QS systems [48]. Several previous studies have shown that halogenated furanones can interact with the LuxR protein and induce conformational changes due to rapid proteolytic degradation of the furanone-LuxR complex, which in turn destabilises the AHL-dependent transcriptional activator [49]. However, independent studies confirmed that halogenated furanones without alkyl chains were strongly toxic to the planktonic cell. While furanones with long alkyl chains were shown not to reduce the biofilm formation [50,51]. The present study confirmed that furanone without alkyl chains has high toxicity against the *E. coli* Top10 pSB1A3-BBaT9002 strain, as it reduced 50% the bacterial growth when dosed at 5 × 10−<sup>5</sup> M. In fact, at 1 × 10−<sup>3</sup> M, indeed, furanone can kill all bacteria, diagnostic that the lack of alkyl chain is at play in the induced toxicity. It has been suggested that the increase of water solubility may explain this effect [51]. Interestingly, when dosed at 5 × 10−<sup>6</sup> M, the results showed that furanone could reduce the fluorescence production at no detrimental expense of the bacterial growth (Figure 2c). A possible explanation to these phenomena might be that some compounds when applied at sub-lethal concentrations will not kill bacterial but delay the QS activity because bacteria become weaker than at the normal condition. This phenomenon gave us the illusion that those compounds were QSIs, but when we increased the dose, we noticed that they had an antibacterial activity. The observed antimicrobial effect at furanone concentrations ≥ 5 × 10−<sup>6</sup> M (Figure S10) prompted us to search QSIs with lower associated toxicity. Closer attention should be paid to this phenomenon to discriminate between potential QSIs more carefully.

Group 2 candidates included compounds with at least one aromatic ring. Vanillin (**5**), proposed as a less toxic alternative to brominated furanones, revealed a relative reduction of the biosensor's fluorescent response of 27.1 % when applied at 1 × 10−<sup>3</sup> M (Figure 3a and Figure S7), an effect that was accompanied by a discrete, 16.0% reduction of cell growth (Figure 3b and Figure S7). Belonging to the same group, the well-reported QSI, *trans*-cinnamaldehyde (**6**) revealed a potent capacity to reduce the QS-based response of the biosensor to up to 68.3% (Figure 3a and Figure S7) when applied at 1 × 10−<sup>3</sup> M, with a slight reduction of cell growth (18.6%; Figure 3b and Figure S7). Surprisingly, related compounds caffeic acid (**7**) and *trans*-anethole (**8**) were apparently innocuous to the *E. coli* biosensor even at concentrations of 1 × 10−<sup>3</sup> M (Figure 3a,b) while capsaicin (**9**) and CAPE (**10**) inhibited cell growth by 12.8 and 24.4%, respectively (Figure 3b).

Vanillin (**5**) is a well-known food flavouring agent and is a major constituent of vanilla pods. Its QS inhibition activity has recently been demonstrated in *Aeromonas hydrophila*, *Agrobacterium tumifaciens* NTL-4, *Chromobacterium violaceum* CV026 when applied at 250 µg/mL (1.64 mM), where it showed significant inhibition in short-chain AHLs (C4-HSL (69%) and 3OC8-HSL (59.8%)), followed by C6-HSL(32%), and C8-HSL (28%), but lower inhibition in long-chain AHLs (C14-HSL (13.5%) and C10-HSL (12%)). It also reduces the biofilm formation on the reverse osmosis membrane of *A hydrophila* [52]. Our own experiments confirmed that vanillin at 1 × 10−<sup>3</sup> M can inhibit the QS activity with 3OC6HSL 27.1%. It has been speculated that the possible mechanism whereby vanillin inhibits QS activity is that it interferes with the binding of the short-chain AHLs to their cognate receptor [52]. Our study offers experimental evidence that vanillin may also interfere with binding of 3OC6HSL to its cognate LuxR receptor.

*trans*-Cinnamaldehyde (**6**) is a component of cinnamon and cassia essential oils, and it is commonly present in food as a flavouring agent and fungicide. In a previous study, it was shown that 200 µM cinnamaldehyde can reduce by 70% the fluorescence intensity due to the expression of GFP, induced by 3-oxo-C6-HSL in a bioreporter *E. coli* ATCC 33,456 pJBA89, and also effective at inhibiting AI-2 mediated QS [53]. Furthermore, Brackman et al. proved *trans*-cinnamaldehyde at concentrations < 1 mM shifts the SDS-PAGE mobility of LuxR-DNA. They concluded that *trans*-cinnamaldehyde and cinnamaldehyde derivatives interfere with AI-2 based QS in various *Vibrio* spp. by decreasing the DNA-binding ability of LuxR [54]. Recent molecular docking analysis studies suggested that *trans*-cinnamaldehyde interacts with LasI and EsaI substrate binding sites thus inducing the QS inhibition activity [55]. Our own studies also confirm that *trans*-cinnamaldehyde inhibits the LuxR-mediated GFP production in the *E. coli* Top 10 biosensor by 68.3% at 1 × 10−<sup>3</sup> M concentration. This strain does not express the LuxI gene, thus it cannot produce LuxI synthase of 3-oxo-C6-HSLs but can constitutively overexpress LuxR protein. In our related study, we have shown that *trans*-cinnamaldehyde not only inhibits the expression of GFP, but it also retards its kinetics [38]. We suggest that there are maybe two mechanisms at play for QS inhibition involving LuxR. Firstly, the three-carbon aliphatic side chain of *trans*-cinnamaldehyde could interfere with the binding of 3OC6HSL to LuxR receptor; secondly, it can also decrease the DNA-binding ability of LuxR dimers, as previously suggested.

*trans*-Cinnamaldehyde (**6**)-related compounds, namely caffeic acid (**7**) and *trans*-anethole (**8**), were also tested on their QS inhibition activity. The results showed that neither of them had QS inhibition activity nor antibacterial effect at the tested concentration of 1 × 10−<sup>3</sup> M. Caffeic acid is a phenolic acid that has various documented beneficial biological properties. Besides being a powerful antioxidant, it also has anticancer, anti-inflammatory and antiviral activities [56]. *trans*-Anethole, in turn, is also a natural component of anise seeds and fruits and it is used as a flavouring ingredient. Until now, its antibacterial and QS inhibitory activities have not been tested.

Comparing the chemical structures of caffeic acid and *trans*-anethole with that of *trans*-cinnamaldehyde, we can clearly see that neither the catechol phenolic nor the aromatic ester groups of caffeic acid and trans-anethole, respectively, have any effect on increasing the binding affinity with the LuxR protein. These results may reflect the importance of the unsubstituted aromatic ring in *trans*-cinnamaldehyde to make π-π interactions with the receptor residues at the binding site of LuxR. In this regard, the phenolic residues of caffeic acid can act as the H-bond donors, but not acceptors, at the hydrogen bond acceptor binding domain of the binding pocket of LuxR, hence, decreasing its overall binding efficiency. Similarly, in trans-anethole, the presence of the methyl ester substituent at the aromatic ring may be is enough to prevent its efficient binding with the binding site of LuxR. However, vanillin, with one phenolic, one methyl ester and an aldehyde substituent at the aromatic ring, does show QS inhibition activity as discussed above. Therefore, our study is consistent with the idea that the role of the presence of H-bond acceptor groups along with the the π-π interactions of the aromatic ring, is what determines the overall affinity of compounds to bind with LuxR receptor.

Capsaicin (**9**), structurally related to vanillin (**5**), is a natural alkaloid extracted from fruit of Capsicum family and it is responsible for the pungency of chili peppers. Its structure has an aromatic ring and a long hydrophobic aliphatic chain with a polar amide group. Most studies with capsaicin have addressed its pharmacology and clinical applications [57]. Only a few studies have focused on the inhibition activity of the growth of the gastric pathogen *Helicobacter pylori* [58,59]. One such study found that capsaicin did not inhibit the growth of a human fecal commensal *E. coli* strain [58]. So far, the QS inhibitory activity of capsaicin has not been documented. Our results show that even at the concentration of 1 × 10−<sup>3</sup> M, capsaicin did not significantly reduce the QS activity of the *E. coli* Top10 pSB1A3-BBaT9002 biosensor, and it decreased the bacterial growth only by 12.8% (*p* < 0.05). If we compare the structures of capsaicin with that of vanillin, we can observe that they share identical substitution positions in their aromatic ring, except that vanillin has not the long aliphatic chain, but only a strong aldehyde H-bond acceptor group. Our results indicate that the increase in the length of the aliphatic chain does not lead to an increase in binding affinity. CAPE (**10**) is the main active component of propolis extract. Its chemical structure is described as the ester of caffeic acid and phenetyl alcohol; hence, it is a catechol ring with an ester chain bearing another aromatic ring. CAPE has known bioactivities such as antimicrobial, anti-inflammatory and cytotoxic activities. About the antimicrobial activity of CAPE, it has been reported that can inhibit the 60% *E. coli* growth when the concentration is over 60.6 µM. This has been explained as the result of the synthesis of reactive oxygen species that damage the outer membrane of the bacteria-induced by CAPE [60,61]. Our experiments contrast with this study, as we found that CAPE only inhibits the growth of the *E. coli* Top10 pSB1A3-BBaT9002 strain by 24.4% when applied at 1 × 10−<sup>3</sup> M. As a possible explanation for the observed discrepancies between the results of our study and the previous ones may stem in the distinct protocols of the assays to quantify the bacterial growth rate. Overall, no QS inhibition activity was shown on the surviving bacteria after treatment with CAPE.

Group 3, comprised by heterocyclic compounds, revealed candidates with important QSI capacity, namely caffeine (**11**) and genipin (**12**), which have been shown to reduce the QS-based response of the biosensor by 47% and by 43.3% when applied at 1x10−<sup>3</sup> M and 5x10−<sup>5</sup> M, respectively (Figure 3a,c, Figures S7 and S8) and with negligible effects on cell growth (Figure 3b–d, Figures S7 and S8). Belonging to the same group, gardenoside (**13**) did not show any apparent effect on the *E. coli* biosensor at the doses tested (Figure 3a,b).

Caffeine (**11**) is yet another alkaloid which occurs in coffee cherry and tea. Documented uses of caffeine include as a pesticide to kill certain larvae, insects and beetles. Some studies have shown that caffeine, applied at 2.5 mg/mL (≈12.8 mM), can retard the growth of *E. coli*, *Enterobacter aerogenes, Proteus vulgaris*, and *P. aeruginosa* within a short time. The first time caffeine was tested as QS inhibition compound against *C. violaceum* CV026 and *P. aeruginosa* PA01 strains, it was found that when applied at 0.3–1.0 mg/mL (≈1.5–5.0 mM), it inhibits violacein production in *C violaceum* CV026, and short chain AHLs production and swarming motility in *P. aeruginosa* PA01 [62]. Our experiments also proved that caffeine, applied at 1 × 10−<sup>3</sup> M to the *E. coli* Top10 pSB1A3-BBaT9002 biosensor strain, it can inhibit 47% GFP production without affecting bacterial growth. Our results, along with previous studies [62], suggest that caffeine has a broad spectrum of QS inhibition activities in different bacterial species; interestingly, all share in common to contain the AHL-regulated QS system. As earlier reported by Norizan et al. Caffeine did not degrade C6-HSL [62], so we hypothesise here that it can be a competitor that binds with AHL receptors because the keto groups from the aromatic ring can also form strong hydrogen bonds with type I QS LuxI/LuxR receptors.

Also in Group 3, genipin (**12**) is an iridoid compound isolated from *Gardenia jasminodies* Ellis fruits. It is the aglycone derivative of geniposide. It was initially identified as a protein cross-linking agent but can also inhibit the production of nitric oxide by downregulating the activity of nuclear factor-κB (NF-κB) [63]. It is also a cell-permeable inhibitor with anti-inflammatory and anti-angiogenic activity mediated by the induction of apoptosis in hepatoma and hepatocarcinoma cell lines [64]. A number of studies have also shown that genipin cross-links chitosan nano- and microparticles, thus allowing them to be used for QS inhibition and the delivery of antimicrobial drugs [65,66]. We found, for the first time, that genipin on its own can inhibit GFP production in the *E. coli* Top10 pSB1A3-BBaT9002 biosensor, without exhibiting significant toxicity, more effectively than a diverse spectrum of alternative compounds. Knowing that genipin possesses inherent fluorogenic properties, we wondered whether the observed effect on the fluorescent response of our biosensor could be due to an artefact associated with the direct fluorescence quenching [67]. We performed a series of extra experiments where we tested genipin on the control *E. coli* strain Top10 pBCA9445-jtk28282::sfGFP strain (Supporting Information). We found that genipin concentrations ranging from 9.95 × 10−<sup>10</sup> to 1.19 × 10−<sup>4</sup> M did not exert significant fluorescence quenching on the control strain (Figures S2 and S3). Nevertheless, the fact that the fluorescence over growth profiles of both bacterial strains widely differ (Figure S1) make these comparisons a difficult task. Moreover, both strains express different GFP species (namely GFPmut3b and sfGFP in the biosensor and control strain, respectively) and their expression vectors widely diverge (see Supporting Information) [68,69]. Future efforts should be focused on validating the effects of genipin by using a control *E. coli* Top10 strain expressing GFPmut3b constitutively from a modified version of the pSB1A3-BbaT9002 plasmid. On our hands, genipin and related compounds are promising candidates for the development of novel therapeutic approaches based on the inhibition of QS in bacteria. Our data showed a significant difference between the QS inhibition activities of genipin and the closely related glycosylated iridoid compound, gardenoside. These two iridoids share similar structures, differing only with respect to the glycosylation in the hydroxyl group of C1, and in the hydroxyl group at position C8 of the heterocyclic structure, in gardenoside. This result may have a biological significance in host-pathogen interactions via inhibition of QS, as glucose oxidase (GOD) is synthesised by many plants, fungi and bacteria. This hypothesis would need further experimental validation.

Quinolone- and phenazine- like compounds belonging to Group 4 (Table 1) are reported to play important roles in QS-regulated phenotypes of *Pseudomonas spp* [70]. These comprise the alkylquinolone PQS (**14**) signal of *P. aeruginosa* and three structurally-related synthetic quinolones, namely MOQ (**15**), HOQ (**16**) and MHOQ (**17**) (Table 1); bearing a bicyclic core structure with different substituents. Among these three structurally-related synthetic compounds, only MOQ demonstrated significant relative QS inhibition activity of 62.2%. The rest of tested compounds, namely HOQ and MHOQ, did not have any detectable activity, neither QS inhibition nor antibacterial (Figure 3c,d and Figure S9). Also, compound PQS comprised not only one six-carbon ring but also a long carbon chain at the six-member heterocyclic ring, which showed an important QSI activity, by inhibiting the QS-based response of the biosensor by 43.1%, with a negligible effect on cell growth (Figure 3c,d and Figure S8). Compounds PYO (**18**), PCA (**19**) and PMS (**20**) are similarly with three rings, but only PYO and PMS have high GFP inhibition and high toxicity, compound PCA also has QQ activity, inhibiting GFP production by 33.7%, but no toxicity (Figure 3c,d).

Group 4, comprised heterocyclic compounds, some of which can be produced naturally, such as the *Pseudomonas* spp. metabolites (PQS, PCA, PYO), while the rest are synthetic [70]. It is well known that phenazines have antimicrobial activity [71]. Moreover, some of these compounds are known to act as signalling molecules that regulate the QS systems in *P. aeruginosa* as discussed in detail below.

Compound PQS has been found as a QS signal that participates in the *P. aeruginosa* QS network and acts as a link between las and rhl quorum sensing, which either directly or indirectly, mediates the expression of 182 genes in *P. aeruginosa* [72,73]. Besides the intraspecific signalling role of PQS in *P. aeruginosa* involving interactions with its cognate LasR-like PqsR receptor and non-cognate-LuxR-like-RhlI, there is no evidence to the date on PQS interference with other LuxR-based QS regulation circuitries. Despite the fact that there is currently no crystal structure available for RhlI, it is known that it shows significant sequence divergence from TraR, our prototype for docking studies. Thus, it is unknown how PQS binds to RhlI in *P. aeruginosa*. A recent paper from Mukherjee et al. explored ligand-receptor binding of PqsR with C4HSL, by generating a homology model based on the *E. coli* SdiA structure, which is a LuxR-like closest homolog of PqsR [74]. In SdiA and other LuxR-type

proteins, the highly conserved amino acids Trp68 and Asp81 interact with the amide group-oxygen and the amide group-nitrogen, respectively, of the cognate C4HSL. Other conserved residues, such as Tyr72 and Trp96, are required for hydrophobic and van der Waals interactions with the ligands [74]. These interactions seem to correlate with our docking predictions of OOHL establishing H-bonding with Trp57 and hydrophobic interactions with Tyr53 and Trp85 in the binding site of TraR (Figure 5a and Table 1). A similar scheme of PQS predicted H-bonding and hydrophobic interactions with Trp57 and Tyr61 (Figure 5c,d and Table 1) would in principle serve as a rationale for a strong affinity of PQS to LuxR homologs and the observed quorum quenching effect on the biosensor's LuxR-regulated response (Figure 3c and Figure S8).

Phenazines are a well-known family of pigments that are secreted from *P. aeruginosa*. Among phenazines, PYO is widely known due to its cytotoxic and redox activities [75]. Importantly, PYO is a terminal signalling factor in the QS network of *P. aeruginosa* [76]. PYO is also an intercellular signal that triggers specific responses in enteric *E. coli* and *Salmonella enterica*, via the SoxR regulon. Whether this kind of signal transduction is also involved in our *E. coli* biosensor in the presence of PYO is a question that we cannot elucidate at present [77]. Also, it is well known that PYO interacts with molecular oxygen to form ROS species that change the redox balance of the cells and that GFP fluorescence can be affected by the presence of ROS species [78,79]. To shed some light on extra effects of PYO on fluorescence quenching and/or the metabolism of *E. coli* Top 10 cells, we decided to perform extra experiments applying increasing PYO concentrations on both the *E. coli* Top10 pSB1A3-T9002 (Figure S4) biosensor and the *luxR*-deficient *E. coli* Top10 pBCA9445-jtk28282::sfGFP (Figures S5 and S6). Strikingly, we found a lack of dose-response effect of PYO on the fluorescence of both strains (Figures S4–S6), together with strong variability among experiments (cf. Figures S5 and S6). These disparate results could arise from some of the transductory and/or redox activity of PYO on our *E. coli* Top10 cells and need further investigation.

PCA (**19**) is yet another redox-active phenazine pigment that is produced from *P. aeruginosa* [80]. PCA is known to be a broad-spectrum activity compound that inhibits the growth of several plant pathogenic species (e.g., *Corynebacterium fascians, Agrobacterium tumefaciens, Erwinia aroideae, Diplodia zeae*) [81,82]. Further studies have found that PCA is precursor for more complex phenazine metabolites, such as 1-hydroxyphenazine, phenazine-1-carboxamide and PYO [76]. Yun Wang et al. found that PCA may shift the redox equilibrium between Fe(III) and Fe(II) in *P. aeruginosa* [83]. Even though PCA and PYO, both are redox-active phenazines, in our study, we found that, at 5 × 10−<sup>5</sup> M, PYO has strong toxicity to *E. coli* Top 10 pSB1A3-BBaT9002 strain, while PCA only inhibits the fluorescence intensity but has no effect on bacterial growth. The different effect of PYO and PCA can be attributed to the fact that PCA may help the *E. coli* alleviate Fe(III) limitation by reducing it to ferrous iron [Fe(II)], thus promoting the bacterial growth [83], thus allowing to observe the QS inhibition activity. We will discuss further this aspect in the context of the in silico molecular docking results below.

PMS is a simple phenazonium salt and an electron acceptor and carrier in biochemical oxidation/reduction studies [84,85]. It is also a O<sup>2</sup> <sup>−</sup> generating agent that can increase intracellular H2O<sup>2</sup> levels and lead to the formation of free radicals that can affect bacterial growth [85]. This explains why in our experiments PMS shows strong toxic effect against *E. coli* Top 10 pSB1A3-BBaT9002 strain. As explained above, this strain is lacking the SoxR regulon to confer resistance against redox stress.

Previous studies have proved that phenazines are antibiotic compounds that can inhibit microbial growth because of the redox-active effect. From our own studies, it can be argued that when phenazines (namely, PCA, PYO, PMS) are applied at the same concentration of 5 × 10−<sup>5</sup> M, only PYO and PMS are toxic to the *E. coli* Top10 pSB1A3-BBaT9002 strain, as we mentioned above. Interestingly, PCA, does not inhibit bacterial growth but has QS inhibition activity. Several studies have also noticed this phenomenon. Morales et al. found that lower concentrations of PCA, PYO and PMS inhibited the fungal yeast-to-filament transition and affected the development of *C. albicans* wrinkled colony biofilms but allowed growth. However, those phenazines have anti-candida activity when the concentration is higher than 500 µM, which means that those compounds have different

biological effects at different concentrations [86]. Furthermore, Skindersoe et al. used sub-inhibitory concentrations of antibiotics in *P. aeruginosa* and found that lower doses of antibiotics could modulate gene expression, so that they interfere with QS signalling [87]. To the best of our knowledge, this dual concentration-dependent activity of phenazines had not been reported before to operate in a mutant *E. coli* Top10 pSB1A3-BBaT9002 strain. However, recently, it has been hypothesised that it is a general mechanism of action of many compounds [88].

For the identification of functional group and their arrangement in the binding site required for binding ligand, GRID map was generated by using three different chemical probes i.e., H bond donor (HD), H bond acceptor (HA) and DRY probe. Grid-MIFs generated for TraR indicated (Figure 4) that acceptor interaction points and hydrophobic patches are dominant in comparison to donor interactions points at the binding. Comparing the functional group present in ligands with the GRID-MIFs (Figure 4) it is clear that because of big hydrophobic patch in the center of cavity, hydrophobic interaction from carbon either from aromatic ring or long chain carbon make very favorable interaction. Apart this one donor group at one side of aromatic ring makes favorable interaction with the Trp 57. Hydrogen bond interaction with Trp 57 is identified as important interaction in receptor substrate interaction. Apart from HOQ we found this interaction in all other five ligands. Whereas at the other side of ring one donor or one acceptor would also make favorable interaction with the Thr129 or water.

Consequently, we identified that in the same ringside two functional groups e.g. an OH group just next to the O (acceptor group), do not make a favourable interacting group. This can be observed in the docking score results for HOQ and MHOQ both with lower scores than MOQ. This might explain that in vitro HOQ and MHOQ, did not exhibit QS inhibition activity, while MOQ, that lacks the 3-hydroxyl group did exhibit QS inhibition activity experimentally. PQS showed better score, i.e., PQS-conf A (−8.04) and PQS-conf B (−6.59), in comparison to HOQ and MHOQ because of the long alkyl chain and an overall more favourable hydrophobic interaction (Table 1). In the PQS-conf A (−8.04), the H-bond between the Trp 57 and ligand is missing in this conformation, but because of PQS has a longer alkyl chain than OOHL, a H-bond with Thr 129 and putative H-bond with water, PQS-conf A has a better score than the other compounds. Whereas in PQS-conf B (−6.59) the H-bond is present between ligand O and Trp 57, but because of the OH just next to O is not a favourable interaction (according to GRID-MIFs), hence its lower score. This analysis clearly indicates that OH next to O is not a favourable functional group for interaction in ligand PQS, which affects the pose and docking score. Moreover, because of the H-bond interactions and more hydrophobic interactions in comparison to natural ligand OOHL, this compound is better over other ligands. Along with PQS, PCA has also shown a high dock score compared to other ligands. We argue that this is the result of the combined effect of the three aromatic rings making more hydrophobic interaction, the deprotonated N present in the aromatic group involved in H-bond interaction with Trp57, and the carboxylate function group involved in H-bond interaction with Thr129 and also probably with water. These favourable interactions might explain the high QS inhibition activity observed for PQS and PCA.

Finally, Group 5 contained a more diverse collection of organic molecules with complex structures. Experimentally, with the exception of compounds itaconic acid at 1 × 10−<sup>3</sup> M and berberine at 5 × 10−<sup>5</sup> M, none of these compounds showed QS inhibition activity. However, we found that polygodial exhibited strong bacterial growth inhibition. Polygodial is a bicyclic sesquiterpene dialdehyde, isolated from different traditional medicinal plants such as *Polygonum hydropiper* and *P. punctatum* [89]. Kubo et al. showed that polygodial has the antibacterial activity against various bacteria, not only as a surfactant to form the pyrrole with primary amine groups at the plasma membrane, thereby disturbing the balance of the membranes, but also may react with various intracellular components when it enters into the cells after the membrane damaged [89]. We also proved that polygodial has high antibacterial activity that suppressed almost completely the growth of *E. coli* Top10 pSB1A3-BBaT9002 strain when dosed at 1 × 10−<sup>3</sup> M.

Berberine is an isoquinoline-type alkaloid isolated from *Coptidis rhizomaand* ("huang lian" in Chinese), a plant used in traditional Chinese medicine, and from other plants. It has been reported that when the concentration is at 30-45 µg/mL could exhibit an antibacterial effect and inhibit biofilm formation of *Staphylococcus epidermidis*. Whether the biofilm formation inhibition of berberine observed in Gram-positive bacteria is connected with the QS regulation is not confirmed [90]. However, recent studies have shown that berberine inhibits the QS in Gram-negative bacteria including antimicrobial-resistant *E. coli* strains, *P. aeruginosa PA01*, *C. violaceum* and *Salmonella enterica* [91,92]. Sun et al. investigated the QS inhibition activity of berberine in antimicrobial-resistant (AMR) *E. coli* strains and found that berberine inhibited biofilm formation and downregulated QS-related genes *luxS*, *pfS, hflX*, *ftsQ*, and *ftsE* of AMR *E. coli* strains at 1/2 (640 µg/mL) or 1/4 (320 µg/mL) minimal inhibitory concentration (MIC) [90]. Thus the tested berberine concentrations of berberine were tested by Sun et al. were ≥ 9.5 times higher than ours (cf. ≥ 160 µg/mL and 16.8 µg/mL in Sun et al.'s and our study, respectively). Moreover, the AMR *E. coli* QS system is a LuxS/AI-mediated system, unrelated to the LuxR-based circuitry present in our biosensor. Under our setting, we found no QS inhibition at a berberine concentration of 50µM (16.8 µg/mL). The lower concentration tested in our assays may explain the observed lack of QS inhibition activity. We decided to limit berberine concentration to 50 µM due to solubility problems at higher concentrations. Further efforts should be focused on testing the QS inhibition potential of berberine and other related compounds at concentrations comparable to those of Sun et al.'s and exploring whether the strong effect observed on LuxS-based QS systems can be extrapolated to LuxR-regulated circuitries.

#### **4. Materials and Methods**

#### *4.1. Library of Tested Chemical Compounds*

Compounds were selected according to their chemical structure and were divided into five groups. They were in all cases of high purity (≥90%) and were either commercially available or synthesised. They were shipped in glass vials as powders or in liquid form and were dissolved in water or organic solution (ethanol or methanol) before use. The details for each compound are given in Table 2. Each is assigned a reference number used throughout this manuscript. 3-Oxohexanoyl-homoserine lactone (3OC6HSL) and all other chemicals were of analytical grade and, unless otherwise stated, were purchased from Merck KGaA (Darmstadt, Germany).




**Table 2.** *Cont*.

<sup>1</sup> The Group column refers to the classification based on chemical structural features (Figure 1), as explained in the text; <sup>2</sup> Synthesised by Prof. Susane Fetzner according to the method of Eiden et al. [93]. HPLC and UV absorption analysis indicated a purity of over 90%; <sup>3</sup> Synthesised by Prof. Susane Fetzner according to the method of Evans and Eastwood [94]. HPLC and UV absorption analysis indicated a purity of over 90%; <sup>4</sup> Synthesised by Prof. Susane Fetzner according to the method of Cornforth and James [95]. HPLC and UV absorption analysis indicated a purity of over 90%.

#### *4.2. Bacterial Strains*

The QQ activity of the 23 selected compounds was determined using the *E. coli* Top 10 strains listed below. The BioBrick standard biological sequence BBa\_T9002, ligated into vector psb1a3 (http://partsregistry.org/Part:BBa\_T9002), was a gift from Prof. Anderson (UC Berkeley, CA, USA). The sequence BBa\_T9002 was introduced by chemical transformation into *E. coli* Top 10 (Invitrogen, Life Technologies Co., Leicestershire, UK) and single-colony cultures from the transformed strain were stored as 30% glycerol stocks at −80 ◦C as described in Section 2.3 below. The sequence BBa\_T9002

comprised the transcription factor (LuxR), which is constitutively expressed, but it is active only in the presence of the exogenous autoinducer signalling molecule 3OC6HSL. At an adequate concentration, two molecules of 3OC6HSL bind to two molecules of LuxR and activate the expression of GFP (output), under the control of the lux pR promoter from *Vibrio fischeri*. The fluorescence biosensor was calibrated for different 3OC6HSL concentrations, as described in our previous studies [4]. An *E. coli* strain Top10 (Invitrogen, Life Technologies Co., U.K.) was transformed with plasmid pBCA9445-jtk2828, carrying a superfolder version of the *gfp* gene (*sfgfp*), which was kindly donated by Prof. Anderson Lab (UC Berkeley, Berkeley, CA, USA). The transformed strain expresses sfGFP constitutively and was used as control culture to test possible fluorescence quenching artefacts of genipin and PYO that could account for the effects observed in the fluorescence *E. coli* Top10 pSB1A3-BBaT9002 biosensor (Supporting Information).

#### *4.3. Growth Media and Glycerol Stocks Preparation*

Bacterial strains were cultivated using on Luria-Bertani (LB) and M9 minimal medium purchased from BD GmbH (Heidelberg, Germany). We inoculated 10 mL of LB broth supplemented with 200 µg/mL ampicillin with a single colony from a freshly streaked plate of Top10 containing BBa\_T9002 and incubated the culture for 18 h at 37 ◦C, shaking at 100 rpm. Glycerol stocks were prepared as described in our previous studies [38]. Briefly, a 500 µL aliquots of overnight bacterial culture were mixed with 500 µL 30% sterile glycerol in 1.5 mL plastic vials and stored at −80 ◦C. Prior to each experiment, an aliquot of a glycerol stock from the single culture was diluted 1:1000 into 20 mL M9 minimal medium supplemented with 0.2% casamino acids, 1 mM thiamine hydrochloride and 200 µg/mL ampicillin (AppliChem GmbH, city, Germany). The culture was maintained under the same conditions until the OD600 reached ~0.15 (~5 h).

#### *4.4. E. coli Top10 Fluorescent Biosensor Assay*

Each tested compound was dissolved in MilliQ water or 100% organic solution (ethanol or methanol) according to their solubility at a high concentration of 200 mM, then diluted with MilliQ water to produce samples at six concentrations: 2 × 10−<sup>2</sup> , 1 × 10−<sup>2</sup> , 1 × 10−<sup>3</sup> , 1 × 10−<sup>4</sup> , 1 × 10−<sup>7</sup> and 1 × 10−<sup>8</sup> M; however, some compounds can only be prepared at a maximal concentration of 50 µM given by their water solubility. The 3OC6HSL was dissolved in acetonitrile to a stock concentration of 100 mM and stored at –20 ◦C kept in a sealed glass vial. Prior to each experiment, serial dilutions from the AHL stock solution were prepared in water to produce solutions with a concentration ranging from 100 mM to 10 nM. We then mixed 10 µL 3OC6HSL solution with 10 µL of the diluted compounds at different concentrations in the wells of a flat-bottomed 96-well plate (cat. # M3061, Greiner Bio-One, city, state abbrev if USA, country) and each well was then filled with 180 µL aliquots of the bacterial culture to test for QS inhibition activity. The final inhibitor concentrations, therefore, ranged from 1 × 10−<sup>3</sup> M to 5 × 10−<sup>10</sup> M. Several controls were also set up. Blank 1 contained 180 µL of M9 medium and 20 µL of MilliQ water to measure the absorbance background. Blank 2 wells contained 180 µL of bacterial culture and 20 µL of MilliQ water to measure the absorbance background-corrected for the cells. Finally, positive control wells contained 10 µL of water plus 10 µL 3OC6HSL solution and 180 µL of the bacterial culture to measure the fluorescence background. The plates were incubated in a Safire Tecan-F129013 Microplate Reader (Tecan, Crailsheim, Germany) at 37 ◦C and fluorescence measurements were taken automatically using a repeating procedure (λex = 480 nm and λem = 510 nm, 40 µs, 10 flashes, gain 100, top fluorescence), absorbance measurements (OD600) (λ = 600 nm absorbance filter, 10 flashes) and shaking (5 s, orbital shaking, high speed). The interval between measurements was 6 min. For each experiment, the fluorescence intensity (FI) and OD<sup>600</sup> data were corrected by subtracting the values of absorbance and fluorescence backgrounds and expressed as the average for each treatment. Data were presented as FI/OD<sup>600</sup> versus incubation time. All measurements were taken in triplicate.

#### *4.5. Protein Structure File, Ligand Database*

The X-ray crystal structure of *Agrobacterium tumefaciens* TraR was downloaded from the Protein Data Bank (PDB ID 1L3L) and used for computer docking. All the water molecules were removed except one molecule in the binding pocket, which plays an important role in interaction and forms the hydrogen bond with the autoinducer OOHL of TraR protein. To define the grid box of TraR protein, OOHL was used as a ligand to select spheres and also followed with the Information from the previous study [18]. The 2D structures of six compounds (OOHL, MOQ, HOQ, MHOQ, PCA and PQS) were drawn manually using Marvin sketch v6.1.3 (ChemAxon Ltd., Budapest, Hungary) and saved as MDL mol files. The mol files were merged into a single mol file and likewise converted to 3D structures using Discovery Studio 3.5 client software. PyMol was used for visualisation and molecular modelling.

#### *4.6. Molecular Docking Studies*

For the generation of GRID-MIFs (molecular interaction fields) at the TarR binding site where a given chemical group can interact favourably, Autogrid program inbuilt in AutoDockTools 1.5.6. was used. For MIF generation, mainly three probed were applied i.e., DRY probe representing hydrophobic interaction, HA probe to representing H bond acceptor groups, and HD probe to representing H bond donor groups. Docking guided by the grid map was performed using Autodock tool. Fifty conformations were generated for each docked substrate. Binding scores between the ligand and protein was evaluated using the autodock utility autoscorer considering the hydrogen bond forces, electrostatic forces, van der Waals forces, solvation energy and entropy.

#### *4.7. Statistical Analysis*

All the experiments were performed in triplicates to validate reproducibility and the P values were calculated statistically by Student's *t*-test. Values were expressed as mean ±SD. A comparison analysis was performed between tests and control.

#### **5. Conclusions**

In this study, we have screened the QS inhibition activity of a library of 23 structurally different compounds against an *E. coli* Top10 pSB1A3-BBaT9002 reporter of AHL-regulated QS. This library included a selection of natural and synthetic compounds that occur naturally in plants and in bacteria species such as *P. aeruginosa*. We were able to establish cues of structure-function relationships for compounds with QS inhibitory activity (e.g., *trans*-cinnamaldehyde, vanillin, caffeine, PQS, PCA). We showed, for the first time, that genipin and MOQ have QS inhibition activity. We also conducted molecular simulations using GRID-MIFs on a selection of compounds (e.g., MOQ, HOQ, MHOQ). Our results aid in the future rational design of novel QS inhibition compounds. For example, the introduction of a 3-methyl group in MOQ may increase the binding affinity substantially to the TraR receptor and hence the QS inhibition activity. This hypothesis could be validated experimentally in future studies. The results of this study may pave the way to future works aimed to fully realise the potential of QS inhibition as an alternative strategy to overcome antimicrobial resistance and biofilms in clinical and other settings.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/1424-8247/13/9/263/s1.

**Author Contributions:** Conceptualiazation, F.M.G., X.Q., C.V.-S. and R.S.; methodology, X.Q. and C.V.-S.; formal analysis, X.Q. and R.S.; resources, B.P. and C.V.-S.; writing—original draft preparation, X.Q.; writing—review and editing X.Q., B.P., R.S., C.V.-S. and F.M.G.; supervision, F.M.G. and B.P.; project administration, F.M.G.; funding acquisition, F.M.G. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Deutsche Forschungsgemeinschaft, grant number: GRK 1549 and the APC was funded by Ideas: European Research Council: FP7 613931. The authors acknowledge financial support by the Open Access Publication Fund of the Westfälische Wilhelms-Universität Münster, Germany.

**Acknowledgments:** X.Q. was recipient of a fellowship from China Scholarship Council. CVS was supported by a pre-doctoral fellowship of the Xunta de Galicia and by a FPU fellowship of the "Ministerio de Educación y Ciencia" of Spain, by a research fellowship of the DAAD (Germany), and research fellowship of the Fundación Pedro Barrié de la Maza (Spain). We acknowledge support from D.F.G., Germany (Project GRK 1549 International Research Training Group 'Molecular and Cellular GlycoSciences'); the research leading to these results has also received funding from the European Union's Seventh Framework Programme for research, technological development and demonstration under grant agreement no. 613931. We are also indebted to Antje von Schaewen for generous access to the Safire Tecan-F129013 Microplate Reader.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **In Vitro Selective Growth-Inhibitory Activities of Phytochemicals, Synthetic Phytochemical Analogs, and Antibiotics against Diarrheagenic**/**Probiotic Bacteria and Cancer**/**Normal Intestinal Cells**

**Tomas Kudera <sup>1</sup> , Ivo Doskocil <sup>2</sup> , Hana Salmonova <sup>2</sup> , Miloslav Petrtyl <sup>3</sup> , Eva Skrivanova <sup>2</sup> and Ladislav Kokoska 1,\***


Received: 21 July 2020; Accepted: 31 August 2020; Published: 3 September 2020

**Abstract:** A desirable attribute of novel antimicrobial agents for bacterial diarrhea is decreased toxicity toward host intestinal microbiota. In addition, gut dysbiosis is associated with an increased risk of developing intestinal cancer. In this study, the selective growth-inhibitory activities of ten phytochemicals and their synthetic analogs (berberine, bismuth subsalicylate, ferron, 8-hydroxyquinoline, chloroxine, nitroxoline, salicylic acid, sanguinarine, tannic acid, and zinc pyrithione), as well as those of six commercial antibiotics (ceftriaxone, ciprofloxacin, chloramphenicol, metronidazole, tetracycline, and vancomycin) against 21 intestinal pathogenic/probiotic (e.g., *Salmonella* spp. and bifidobacteria) bacterial strains and three intestinal cancer/normal (Caco-2, HT29, and FHs 74 Int) cell lines were examined in vitro using the broth microdilution method and thiazolyl blue tetrazolium bromide assay. Chloroxine, ciprofloxacin, nitroxoline, tetracycline, and zinc pyrithione exhibited the most potent selective growth-inhibitory activity against pathogens, whereas 8-hydroxyquinoline, chloroxine, nitroxoline, sanguinarine, and zinc pyrithione exhibited the highest cytotoxic activity against cancer cells. None of the tested antibiotics were cytotoxic to normal cells, whereas 8-hydroxyquinoline and sanguinarine exhibited selective antiproliferative activity against cancer cells. These findings indicate that 8-hydroxyquinoline alkaloids and metal-pyridine derivative complexes are chemical structures derived from plants with potential bioactive properties in terms of selective antibacterial and anticancer activities against diarrheagenic bacteria and intestinal cancer cells.

**Keywords:** plant compounds; diarrhea; antibacterial; anticancer; selectivity

#### **1. Introduction**

The lack of an effective and safe antimicrobial therapy for diarrheagenic bacterial infections is a global health concern, especially in developing countries for children under the age of five years [1]. Although mortality associated with bacterial diarrhea is low in developed countries, the increased incidence rates of inflammatory bowel disease and colorectal cancer have also been attributed to

gut dysbiosis that can result from chronic intestinal infections [2]. Currently, infectious diarrhea is treated using conventional drugs belonging to various classes of antibiotics, such as ceftriaxone, chloramphenicol, ciprofloxacin, tetracycline, metronidazole, and vancomycin. However, the irrational use of antibiotics, including incorrect dose prescription, has led to the development of drug resistance in several pathogens. Additionally, the applications of conventional antibiotics are limited, especially among children in developing countries, owing to high cost and increased risk of side effects including gut dysbiosis. Therefore, there is a need to identify novel antimicrobial agents for infectious bacterial diarrhea to overcome the limitations of conventional antimicrobial drugs [3].

Over the last decades, plant-derived products have become a mainstay in providing novel chemical scaffolds for the development of anti-infective drugs, and therefore antidiarrheal medicinal plants and their bioactive components can be examined first [4]. However, the therapeutic effect of products derived from antidiarrheal medicinal plants is not necessarily based on their antimicrobial activity against the causative agents as other mechanisms can be considered important, such as antimotility and antisecretory effects, for their useful applications [5]. The chance of finding a new plant-derived compound with promising antibacterial activity could therefore be enhanced by the chemotaxonomic approach by examining such novel medicinal plants that are taxonomically related to the species known to bear specific types of phytochemicals with growth-inhibitory effect on diarrheagenic bacteria [6]. In a recent review paper, Kokoska et al. [7] described the in vitro antimicrobial properties and clinical efficacy of plant-derived compounds with their synthetic analogs that are present in products available in the international market as over-the-counter pharmaceuticals, dietary supplements, and herbal medicines for intestinal infections. Benzylisoquinoline alkaloid berberine (e.g., *Hydrastis canadensis*), simple phenol bismuth subsalicylate, the analog of salicylic acid derived from salicin (*Salix alba*), and polyphenol tannic acid (e.g., *Quercus* spp.) were mentioned as examples of the efficient agents with potent activity against some of the gut bacterial pathogens.

As it has recently been proposed, investigations of new antimicrobial agents should be focused on the identification of structures with lowered toxicity to indigenous intestinal microbiota including probiotic bacterial strains. Since there is a common association between dysbiosis and the use of antibiotics, the agents selectively acting against pathogenic microorganisms can prevent the risk of developing diseases, such as chronic bowel inflammation and intestinal carcinoma [2]. Although the composition of the whole gut microbiota comprises a large number of microorganisms forming a complex ecosystem, a preliminary screening of selective antibacterial activity of newly tested agents could be performed by testing the in vitro susceptibilities of particular representatives of each of the three dominant bacterial phyla that can be found in human intestines, and which probiotic function have been described. Among them, we can recognize the strains such as *Bifidobacterium* spp. (Actinobacteria), *Lactobacillus* spp. (Firmicutes) and *Bacteroides fragilis* (Bacteroidetes) [8,9]. Selective in vitro growth inhibitory effect of a plant-derived compound was for example described in the study of Novakova et al. [10], where anticlostridial effect of 8-hydroxyquinoline (*Microstachys corniculata*) was comparably higher than the activities revealed against different strains of bifidobacteria. Chloroxine, the synthetic 8-hydroxyquinoline derivative, is the antimicrobial agent that has also been used as an oral formulation for infectious diarrhea, and disorders of the intestinal microbiota [11]. In dysbiosis, the increased abundance of *Fusobacterium nucleatum* and *Faecalibacterium prausnitzii* is positively and negatively correlated with the risk of intestinal carcinogenesis, respectively [12]. The repression of gut microbiota enhances the susceptibility of host intestinal cells to diarrheagenic bacterial infection and chemical-induced cytotoxicity. Thus, novel antimicrobial agents should not exhibit cytotoxic activity against normal intestinal cells [13]. For example, quinolone antibiotics were reported to inhibit the growth of both bacterial and eukaryotic cells through the same mechanism and consequently enhance the risk of eliciting cytotoxic response [14,15]. On the other hand, the antiproliferative activity of these agents with selective cytotoxic effects against intestinal cancer cells would be a suitable feature in cases of dysbiosis associated with carcinogenesis. There are limited studies on the anticancer effects of conventional antibiotics. For example, Bourikas et al. [16] reported that ciprofloxacin is potent

to inhibit the proliferation of intestinal cancer cell line HT29 in vitro. On the other hand, a strong in vitro antiproliferative activity of plant-derived compounds against various cancer cell lines has extensively been reported, and some are used as a scaffold for anticancer drugs. For example, the alkaloid camptothecin, extracted from the bark of *Camptotheca acuminata*, is currently used as a cytostatic agent for the treatment of colon cancer [17]. We, therefore, suggest that the phytochemicals with known in vitro growth-inhibitory activity against some of the diarrheagenic bacteria could potentially exhibit selective cytotoxic effects on cancer cells. Amongst them, the anticancer effect has generally been reported for quinoline alkaloids [18]. In addition to those earlier mentioned, benzylisoquinoline alkaloid sanguinarine (*Sanguinaria canadensis*) [7,19], and 8-hydroxyquinoline derivatives ferron [20] and nitroxoline [21] are examples of antimicrobial drugs with potent anticancer activity. Another type of synthetic phytochemical analog with reported antimicrobial and anticancer properties is metal-pyridine derivative complex zinc pyrithione (pyrithione found in *Polyalthia nemoralis*) [7,22].

In this study, we compared the selective antibacterial (diarrheagenic/probiotic strains) and cytotoxic (cancer/normal intestinal cells) activities of phytochemicals (alkaloids and phenolics) and their synthetic analogs with those of antibiotics in vitro. For each test compound, values of minimum inhibitory concentrations (MICs), half-maximal inhibitory concentration (IC50), and 80% inhibitory concentration of proliferation (IC80) were assessed. The means of these values (*x-*MIC, *x-*IC50, and *x*-IC80), each defined for a particular type of strain/cell line, were used for calculation of selectivity index (SI) between activities against normal intestinal cells and diarrheagenic strains (SIa), probiotic and diarrheagenic strains (SIb), and normal and cancer intestinal cells (SIc). The aim was to obtain some of the missing data on particular in vitro activities of the test compound, to assist in identifying phytochemicals with scaffolds that possess a potent combination of bioactivities. They could subsequently be utilized in future chemotaxonomic investigation of antidiarrheal medicinal plants with their bioactive components considered for new chemotherapeutic agents against diarrheal infections and associated intestinal cancer diseases. Our results show that 8-hydroxyquinoline alkaloids and zinc pyrithione possess in vitro selective antibacterial properties against diarrheagenic bacteria comparable to ciprofloxacin and tetracycline, with additional in vitro antiproliferative activity against cancer intestinal cell lines. However, in contrary to antibiotics, these compounds generally possess increased cytotoxicity to normal intestinal cells.

#### **2. Results**

#### *2.1. Antibacterial Activity*

As far as the antibacterial activity of antibiotics against diarrheagenic strains is considered, ciprofloxacin and tetracycline exhibited strong growth-inhibitory effect (*x*-MICs = 2 ± 4 and 4.8 ± 8 µg/mL, respectively), while chloramphenicol and ceftriaxone exhibited moderate growth-inhibitory activities (*x*-MIC = 16.5 ± 34 and 61.8 ± 141 µg/mL, respectively). Regarding the significant degree of variation between MICs of these compounds, particular types of pathogenic strains were highly susceptible while some other species were distinctly more resistant. For example, all gram-negative diarrheagenic bacteria were highly susceptible to ciprofloxacin (MICs = 0.016–0.125 µg/mL) and ceftriaxone (MICs = 0.062–0.5 µg/mL). In contrast, their MICs produced against gram-positive pathogens were comparably higher; for the former ranging from 1 to 16 µg/mL and for the latter in the range of 4–512 µg/mL. At least half of the diarrheagenic bacteria were inhibited at the low MICs (1–4 µg/mL) by chloramphenicol and tetracycline, whereas the variations were particularly caused by the weak activities revealed against *Enterococcus faecalis* (MIC = 128 µg/mL) and *Clostridium perfringens* (32 µg/mL), respectively. At the low concentration (MICs = 0.5 µg/mL), tetracycline also inhibited *Clostridium di*ffi*cile* and *Bacillus cereus.* Although metronidazole and vancomycin were generally inactive against diarrheagenic bacteria (*x*-MIC = 651.4 ± 459 and 512.9 ± 404, respectively), they produced a strong inhibitory effect against both clostridial species tested (MICs = 0.5–8 µg/mL), whereas the former also exhibited strong activity against *Escherichia coli* (0.062 µg/mL). The synthetic

analogs of phytochemicals, namely, zinc pyrithione, nitroxoline, and chloroxine, exhibited strong to moderate growth-inhibitory activity against all diarrheagenic bacteria (*x*-MICs = 7.1 ± 4, 12 ± 10, and 24 ± 19 µg/mL, respectively). The activities of these compounds against particular pathogenic bacteria did not have significant differences, however, some of the strains were comparably more susceptible. Regarding that, *B. cereus*, *E. coli*, *Shigella flexneri*, and *Vibrio parahaemolyticus* were highly susceptible to zinc pyrithione (MICs = 1–4 µg/mL). The MIC of chloroxine against both *B. cereus* and *C. di*ffi*cile* was 8 µg/mL. Nitroxoline exhibited strong growth-inhibitory activities (MICs = 2–4 µg/mL) against *B. cereus*, clostridial species, *E. coli*, and *S. flexneri*. Although 8-hydroxyquinoline did not produce significant antibacterial activity against the pathogens (*x*-MIC = 224.4 ± 181), its growth-inhibitory activities were strong against *E. faecalis* (MIC = 4 µg/mL) and *Listeria monocytogenes* (MIC = 1 µg/mL).

Subsequently, the growth-inhibitory activities of different test compounds against probiotic strains were evaluated. Probiotic bacteria exhibited high susceptibility to chloramphenicol (*x-*MIC = 6.2 ± 4 µg/mL) and medium susceptibility to tetracycline, nitroxoline, zinc pyrithione, ciprofloxacin, ceftriaxone, sanguinarine, and vancomycin (*x-*MICs = 19.8±19–47.8±78 µg/mL). The MICs (2–4 µg/mL) of both chloramphenicol and vancomycin against bifidobacteria were similarly low, but the latter produced significantly lower activities against lactobacilli (MICs = 64–256 µg/mL) and *B. fragilis* (MIC = 32 µg/mL). With the exception of *Bifidobacterium breve* and *Bifidobacterium longum* ssp. *longum* (MICs = 8–32 µg/mL), the remaining bifidobacteria were also susceptible to ceftriaxone (MICs = 1–4 µg/mL). However, considering the MIC range (MICs = 0.5–32 µg/mL) shown against lactobacilli and the low activity against *B. fragilis* (MIC = 128 µg/mL), the variation of activities of this drug against probiotic bacteria is quite high. The exceptionally strong activities against *B. fragilis* were revealed by metronidazole and tetracycline (MICs = 0.5 µg/mL), whereas the MIC (4 µg/mL) of nitroxoline against this bacterium was the same as in the case of chloramphenicol. In general, berberine (MICs ≥ 32 µg/mL), ferron (MICs ≥ 64 µg/mL), and phenolic compounds (MICs ≥ 64 µg/mL) did not exhibit significant antibacterial activities against any of the 21 strains. The complete data on growth-inhibitory activities of test compounds against diarrheagenic and probiotic strains, including calculated mean values (*x-*MIC) are presented in Table 1.


**Table 1.**In vitro selective inhibitory activities of phytochemicals, their synthetic analogs, and antibiotics against intestinal bacteria and cells.


**Table 1.** *Cont.*

MIC: minimum inhibitory concentration; IC50: half maximal inhibitory concentration; IC80: 80% inhibitory concentration of proliferation; SD. standard deviation. a Not active (MIC/IC50/80 > 512 µg/mL, the value 1024 µg/mL was used for average calculation). BR: berberine, SG: sanguinarine, 8HQ: 8-hydroxyquinoline, CLX: chloroxine, NXL: nitroxoline, FRN: ferron, ZP: zinc pyrithione, SA: salicylic acid, BS: bismuth subsalicylate, TA: tannic acid, CF: ceftriaxone, CP: ciprofloxacin, MA: metronidazole, VM: vancomycin, CA: chloramphenicol, TC: tetracycline. BC: *Bacillus cereus*, CD: *Clostridium di*ffi*cile*, CP: *Clostridium perfringens*, EF: *Enterococcus faecalis*, EC: *Escherichia coli*, ECS: *E. coli* 0175:H7, LM: *Listeria monocytogenes*, SF: *Shigella flexneri*, SE: *Salmonella* Enteritidis, ST: *Salmonella* Typhimurium, VP: *Vibrio parahaemolyticus*, YE: *Yersinia enterocolitica*, BF: *Bacteroides fragilis*, BA: *Bifidobacterium adolescenti*s, BLC: *Bifidobacterium animalis* spp. *lactis*, BBF: *Bifidobacterium bifidum*, BB: *Bifidobacterium breve*, BL: *Bifidobacterium longum* ssp. *longum*, LC: *Lactobacillus casei*, LR: *Lactobacillus reuteri*, LRM: *Lactobacillus rhamnosus*. *x* -DB: mean MIC for diarrheagenic bacteria, *x* -*PB*: mean MIC for probiotic bacteria, *x -CC*: mean IC50/80 for intestinal cancer cells, FHs 74 Int (intestinal normal cells), SD: standard deviation. SI (Selective Index): (**a**) normal cells/diarrheagenic bacteria, (**b**) probiotic bacteria/diarrheagenic bacteria, (**c**) normal cells/cancer cells.

#### *2.2. Cytotoxic E*ff*ect*

Amongst the test compounds, only alkaloids and related structures exhibited strong cytotoxic activity, while other agents, especially antibiotics, exhibited moderate or no cytotoxic activity. Considering the antiproliferative effect of antibiotics on normal intestinal cells (FHs 74 Int), ceftriaxone, metronidazole, and vancomycin were not cytotoxic at all tested concentrations (IC<sup>50</sup> and IC<sup>80</sup> > 512 µg/mL), whereas tetracycline (IC<sup>50</sup> = 14.7 ± 2.3 µg/mL; IC<sup>80</sup> = 108.2 ± 5 µg/mL), chloramphenicol (IC<sup>50</sup> = 30.7 ± 5.6 µg/mL; IC<sup>80</sup> > 512 µg/mL), and ciprofloxacin (IC<sup>50</sup> = 51.8 ± 27 µg/mL; IC<sup>80</sup> = 129.5 ± 24 µg/mL) were moderately cytotoxic. In case of phytochemicals and their synthetic analogs, salicylic acid (IC<sup>50</sup> = 73.2 ± 4.6 µg/mL; IC<sup>80</sup> = 206.9 ± 69 µg/mL), ferron (IC<sup>50</sup> = 22.6 ± 3.3 µg/mL; IC<sup>80</sup> = 46 ± 2.2 µg/mL), and 8-hydroxyquinoline (IC<sup>50</sup> = 10.7 ± 0.2 µg/mL; IC<sup>80</sup> = 20.3 ± 2.4 µg/mL), revealed moderately cytotoxic effects against FHs 74 Int, whereas the other compounds were cytotoxic (IC<sup>50</sup> values = 0.3 ± 0.1–1 ± 0.1 µg/mL; IC<sup>80</sup> values = 0.5 ± 0.03–26.4 ± 0.8 µg/mL). Considering the antiproliferative effect on cancer intestinal cells, zinc pyrithione, 8-hydroxyquinoline, and sanguinarine were cytotoxic to HT29 (IC<sup>50</sup> values = 0.6, 1.3, and 0.9 µg/mL, respectively) and Caco-2 (IC<sup>50</sup> values = 0.7, 0.3 and 0.8 µg/mL, respectively) cells. Nitroxoline (IC<sup>50</sup> = 1.1 µg/mL) and chloroxine (IC<sup>50</sup> = 1.3 µg/mL) exhibited comparable cytotoxic activity against Caco-2 cells. Zinc pyrithione had the lowest *x-*IC<sup>50</sup> value (0.6 ± 0.05 µg/mL) against cancer cells, followed by 8-hydroxyquinoline (0.8 ± 0.5 µg/mL), sanguinarine (0.8 ± 0.05 µg/mL), nitroxoline (1.8 ± 0.8 µg/mL), and chloroxine (2.5 ± 1.2 µg/mL). Berberine (*x*-IC<sup>50</sup> = 12.2 ± 7 µg/mL), tannic acid (*x*-IC<sup>50</sup> = 31.7 ± 4 µg/mL), and ferron (*x-*IC<sup>50</sup> = 70.8 ± 16 µg/mL), produced moderate cytotoxic activity, while salicylic acid and bismuth subsalicylate (*x-*IC<sup>50</sup> ≥ 253.6 ± 208 µg/mL) did not exhibit significant cytotoxic activity against cancer cells. At relatively high concentrations, some antibiotics exhibited antiproliferative activity against cancer cells, namely: ciprofloxacin, tetracycline, and chloramphenicol (*x-*IC<sup>50</sup> = 100.1 ± 30–355.2 ± 84 µg/mL). The complete data on the antiproliferative activities of test compounds against normal and cancer intestinal cells, including calculated mean values for the latter (*x-*IC<sup>50</sup> and *x-*IC80), are presented in Table 1.

#### *2.3. Selective Toxicity*

The selective antibacterial activities against the pathogens with relatively lower activity against probiotic strains (SIb values range from 0.2–1.1) was revealed by most of the agents exhibiting strong to moderate inhibitory effects on diarrheagenic bacteria, namely: nitroxoline, zinc pyrithione, tetracycline, chloroxine, and ciprofloxacin. In contrast, chloramphenicol and ceftriaxone were more toxic to probiotic strains (SIbs = −0.4 for both). Although the antibacterial activity of berberine, ferron, phenolic compounds, and sanguinarine was in cases of both diarrheagenic and probiotic strains generally insignificant, the results show that these agents were rather toxic to the latter (SIb values range from −1 to −0.03). Due to the minor cytotoxicity revealed against FHs 74 Int, none of the antibiotics exhibited an increased toxicity to normal intestinal cells at the inhibitory concentrations active against diarrheagenic bacteria (SIa values = 0.2–1.8), especially ciprofloxacin and chloramphenicol. In contrast, all of the phytochemicals and their synthetic analogs revealed cytotoxicity to normal intestinal cells at the concentrations they were generally inactive against diarrheagenic bacteria (SIa values range from −1.9 to −0.7). Only 8-hydroxyquinoline (SIc = 1.1) and sanguinarine (SIc = 0.1) exhibited selective antiproliferative activity against cancer cells with the decreased cytotoxic effect on normal intestinal cells. Except these two, other tested compounds were more toxic to normal than to cancer intestinal cells (SIcs = from −1.5 to −0.3), or in the case of ceftriaxone, metronidazole, and vancomycin, they did not show any selectivity (SIcs = 0), as they did not inhibit cell lines at any concentration tested. The data on selective toxicities, including all calculated SI values are presented in Table 1. The curves of in vitro selective concentration-dependent effect of ciprofloxacin, chloroxine, nitroxoline, tetracycline, and zinc pyrithione on the growth of diarrheagenic and probiotic bacteria and of 8-hydroxyquinoline on intestinal normal and cancer cells proliferation are shown in Figure 1.

‐ ‐ *x* ‐ *x* ‐ ‐ **Figure 1.** Selective concentration-dependent effect of chloroxine, ciprofloxacin, nitroxoline, tetracycline, and zinc pyrithione on the growth of diarrheagenic and probiotic bacteria and of 8-hydroxyquinoline on intestinal normal and cancer cells proliferation in vitro. MIC: minimum inhibitory concentration; IC50: half maximal inhibitory concentration; IC80: 80% inhibitory concentration of proliferation. *x* -DB: mean MIC for 12 diarrheagenic bacteria, *x* -PB: mean MIC for 9 probiotic bacteria; Caco-2 and HT29: intestinal cancer cells; Fhs74 Int: intestinal normal cells.

#### *2.4. Principal Component Analysis (PCA)*

The correlation between biological activities and chemical structures of the tested compounds and their groups (antibiotics, phenolic compounds, alkaloids, and related structures) was analyzed using PCA (Figure 2). Although the compounds were distributed equally in all quadrants, detailed analysis revealed specific patterns. The closest correlation was observed in the lower-left quadrant between four antibiotics (ceftriaxone, ciprofloxacin, chloramphenicol, and tetracycline), which indicated that these antibiotics exhibited strong growth-inhibitory activity against diarrheagenic and probiotic strains but did not exhibit distinct cytotoxic activity against normal and cancer intestinal cells. The second highest correlation observed in the lower right quadrant indicated that zinc pyrithione and 8-hydroxyquinolines (excluding ferron) exhibited selective antibacterial activity against diarrheagenic strains along with strong to moderate cytotoxic activity against both types of tested cell lines. The correlation observed in the upper right quadrant indicated that tannic acid, benzylisoquinoline alkaloids, and ferron exhibited moderate to no antipathogenic effect (with negative SIbs) and overall moderate to strong cytotoxic activity. The upper left quadrant contains the remaining antibiotics (metronidazole and vancomycin) and both simple phenols with minimal correlation. These agents exhibited moderate to no growth-inhibiting activity against diarrheagenic strains and practically no cytotoxic activity. Whereas all alkaloids are distributed in lower and upper right quadrants indicating their capability to reveal any type of the tested bioactivities, phenols are spread in the right and left upper quadrants which shows their lack of significant antibacterial activity but a certain degree of cytotoxicity. In contrast, antibiotics are concentrated in the lower-left quadrant slightly overlapping the upper left one, therefore they usually display a significant antibacterial effect which is rarely accompanied by cytotoxicity. ‐ ‐ ‐ ‐ ‐

‐ **‐‐‐‐‐‐‐ ●** ‐ ▲ **∆ □** ‐ **■** ‐ **Figure 2.** Principal component analysis of antibacterial and cytotoxic activities of phytochemicals, their synthetic analogues, and antibiotics against intestinal bacteria and cells in vitro. BR: berberine, SG: sanguinarine, 8HQ: 8-hydroxyquinoline, CLX: chloroxine, NXL: nitroxoline, FRN: ferron, ZP: zinc pyrithione, SA: salicylic acid, BS: bismuth subsalicylate, TA: tannic acid, CF: ceftriaxone, CP: ciprofloxacin, MA: metronidazole, VM: vancomycin, CA: chloramphenicol, TC: tetracycline. Antibiotics [**——-**, •]; Phenolic compounds [ . . . . . . **.** , poly- (N), simple (**∆**)]; Alkaloids and related structures [**\_\_\_\_\_** , benzylisoquinolines (), 8-hydroxyquinolines (), metal-pyridine derivative complex (**\***).

#### **3. Discussion**

The in vitro growth-inhibitory properties of the tested antibiotics have previously been reported for a number of diarrheagenic and probiotic bacteria. However, the bacterial strains tested as well as the methods with criteria used for antimicrobial activity assessment vary frequently among the previous studies. Moreover, the studies reporting the in vitro susceptibilities of probiotic bacteria deal more with clinical isolates [23], and less with standard strains [24]. The present study, therefore, provides the data on in vitro selective antibacterial activities of these antibiotics that can be fairly compared with the same data obtained for phytochemicals and their synthetic analogs. We suggest that the reason behind the increased resistance of probiotic strains differ for particular compounds that showed a selective antipathogenic effect. The growth-inhibitory activities of fluoroquinolones against Gram-positive bacteria are reported to be lower than those against Gram-negative bacteria [15]. Consistent with this finding, bifidobacteria and lactobacilli (Gram-positive) were generally less susceptible to ciprofloxacin than Gram-negative diarrheagenic bacteria that predominate over Gram-positive pathogens in this study. The decreased susceptibility of bifidobacteria to tetracycline might be caused by the presence of specific antibiotic resistance genes [25]. Similar to other third-generation cephalosporins [26], the growth-inhibitory activity of ceftriaxone against Gram-negative bacteria was higher than that against Gram-positive bacteria. However, as a result of significant resistance of the tested Gram-positive pathogens and susceptibility of bifidobacteria, ceftriaxone showed increased toxicity to probiotic strains. The growth-inhibitory activities of some alkaloids and related structures were comparable with those of antibiotics. The antibacterial activity of 8-hydroxyquinoline alkaloids is mediated through the chelation of metals that function as co-factors in various enzymes, which results in the inhibition of RNA synthesis. We suggest that probiotic strains (mainly bifidobacteria) are more resistant to 8-hydroxyquinolines as they are able to sequester iron from the environment [10]. The selective antibacterial activity of 8-hydroxyquinoline against diarrheagenic pathogens seems to be enhanced with chlorine halogenation or by the presence of a nitro group and decreased with iodine halogenation and the presence of a sulfo group, as respectively observed for chloroxine, nitroxoline, and ferron in our study. The in vitro selective anticlostridial effect of 8-hydroxyquinoline with increased resistance of bifidobacteria was previously described in studies of Novakova et al. [10,27,28], Skrivanova et al. [29], and Kim et al. [30]. However, data on its in vitro growth-inhibitory effects against a broader selection of diarrheagenic bacteria are limited. The present study also provides new data on in vitro antibacterial activities of chloroxine against diarrheagenic bacteria in addition to those previously published [31,32]. It has been reported that Endiaron, a chloroxine-containing antimicrobial product used for infectious diarrhea, exhibits antimicrobial activity against the pathogens and does not affect the host indigenous microbiota [11], which is in agreement with the increased resistance of probiotic bacteria described in the present study. Interestingly, the antipathogenic activity of nitroxoline, used to treat urinary tract infections, was higher than that of chloroxine. However, the antibacterial selectivity of nitroxoline against diarrheagenic strains was lower. Out of the intestinal bacteria tested herein, there are only data on in vitro inhibitory effects of nitroxoline against *E. coli* and *E. faecalis* that have been reported before [33]. In spite of zinc pyrithione being only used topically for dermatological infections [7], in relation to the plant compounds and their synthetic analogs in this study, it exhibited the highest growth-inhibitory activity against diarrheagenic bacteria with lowered toxicity to probiotic bacteria. According to our best knowledge, this is the first report on in vitro selective antibacterial activities of zinc pyrithione on intestinal diarrheagenic and probiotic bacteria. Although there is limited knowledge on the mechanism underlying the antibacterial activity of zinc pyrithione, the mechanism may be similar to that of 8-hydroxyquinolines [34]. The weak antimicrobial activities of phenols against diarrheagenic strains are consistent with those reported in previous studies. The effectiveness of phenols in infectious diarrhea may be based on other mechanisms, such as astringent, mucosa-protective, and anti-inflammatory properties, or inhibition of pathogenic enterotoxins [7]. Although clinical studies on extensively used phytochemical berberine have reported comparably higher efficiency than certain antibiotics (e.g., chloramphenicol) [35], our results did not show its significant in vitro

antibacterial activity. A possible reason for this discordance is that berberine rather neutralizes diarrheagenic action of bacteria by inhibiting their enterotoxins, as described by Sack and Froelich [36]. The MICs of both benzylisoquinoline alkaloids against some diarrheagenic bacteria reported in this study were higher than those reported in previous studies. This may be because the inoculum density used in this study was higher than that used in previous studies [7,37].

The mechanism underlying the antiproliferative activity of some antibiotics, such as ciprofloxacin, tetracycline, and chloramphenicol, may be similar to that underlying antimicrobial activity [14,38,39]. Previous studies have evaluated the antiproliferative activity of chemicals derived from ciprofloxacin and tetracycline against cancer cells and suggested their applications in cancer therapy [14,40]. Consistent with the results of this study, previous studies have revealed that other antibiotics are not cytotoxic to eukaryotic cells [41–43]. The antitumor activities of some plant compounds and their synthetic analogs have been investigated previously. In the case of 8-hydroxyquinoline and its derivatives, the interaction with metal ions, namely copper and iron, and their transportation into cells has been reported as crucial for its antiproliferative activity [44]. Freitas et al. [45] reported that 8-hydroxyquinoline derivatives with potent anticancer potential often contain halogen substituents. However, in the present study, nitroxoline exhibited stronger antiproliferative activity against cancer cells than chloroxine and ferron. Previous studies have reported that the underlying mechanism of antitumor activity of zinc pyrithione and sanguinarine involves the inhibition of proteasomal deubiquitinases and microtubule depolymerization, respectively [19,22]. All of the above-mentioned alkaloids and related structures revealed increased toxicity to normal intestinal cells in comparison with their antipathogenic effect, which limits their applications in treating bacterial diarrhea. Although there are no studies reporting oral toxicity or toxicity to the digestive system from berberine, chloroxine, and nitroxoline, we suggest that their safety profile should be further examined and the potential protective role of indigenous gut microbiota against these cytotoxic chemicals should be more deeply studied. Zinc pyrithione, 8-hydroxyquinoline, and sanguinarine are not part of any product intended for internal use. Hence, their dose-dependent toxicological profile and oral safety must be carefully elucidated before any consideration for their application for treating infectious diarrhea associated with intestinal cancer [46].

#### **4. Materials and Methods**

#### *4.1. Chemicals*

Phytochemicals (berberine chloride, 8-hydroxyquinoline, salicylic acid, tannic acid, and sanguinarine chloride) and their synthetic analogs [chloroxine (5,7-dichloroquinolin-8-ol), nitroxoline (5-nitroquinolin-8-ol), ferron (7-iodo-8-hydroxyquinoline-5-sulfonic acid), bismuth subsalicylate, and zinc pyrithione], as well as antibiotics (ceftriaxone sodium, ciprofloxacin, chloramphenicol, metronidazole, tetracycline, and vancomycin hydrochloride), used in this study were purchased from Sigma-Aldrich (Prague, Czech Republic). Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, Prague, Czech Republic) was used to prepare the stock solutions of all test compounds, except those of metronidazole, salicylic acid, vancomycin, and zinc pyrithione, which were prepared using distilled water. Stock solutions of chloramphenicol, tannic acid, and tetracycline were prepared using 96% ethanol (Sigma-Aldrich, Prague, Czech Republic). The chemical structures of individual compounds tested are shown in Figure 3.

‐ **Figure 3.** The chemical structures of the tested phytochemicals, their synthetic analogs, and antidiarrheal antibiotics. **1**. berberine chloride, **2**. sanguinarine chloride, **3**. 8-hydroxyquinoline, **4**. chloroxine, **5**. nitroxoline, **6**. ferron, **7**. zinc pyrithione, **8**. salicylic acid, **9**. bismuth subsalicylate, **10**. tannic acid, **11**. ceftriaxone sodium, 12. ciprofloxacin, **13**. metronidazole, **14**. vancomycin hydrochloride, **15**. chloramphenicol, **16**. tetracycline.

#### *4.2. Bacterial Strains and Growth Media*

‐ ‐ The intestinal bacterial type strains were obtained from the American Type Culture Collection (ATCC, Rockville, MD, USA), Czech Collection of Microorganisms (CCM, Brno, Czech Republic), German Collection of Microorganisms and Cell Cultures (DSMZ, Braunschweig, Germany), and National Collection of Type Cultures (NCTC, London, UK). In accordance with the diversity of diarrheagenic Gram-positive and Gram-negative bacteria responsible for globally distributed

foodborne, waterborne, and nosocomial infections [3,47], the following 12 strains were used in this study: *B. cereus* (ATCC 14579), *C. di*ffi*cile* (DSMZ 12056), *C. perfringens* (DSMZ 11778), *E. faecalis* (ATCC 29212), *E. coli* (ATCC 25922), *E. coli* 0175:H7 (NCTC 12900), *L. monocytogenes* (ATCC 7644), *S. flexneri* (ATCC 12022), *Salmonella enterica* ssp. *enterica* serovar Enteritidis (ATCC 13076), *S. enterica* ssp. *enterica* serovar Typhimurium (ATCC 14028), *V. parahaemolyticus* (ATCC 17802), and *Yersinia enterocolitica* (ATCC 9610). The following nine bacterial strains, which belong to three predominant bacterial phyla in the human gut and exhibit probiotic functions [8,9], were used in this study: *Bacteroides fragilis* (ATCC 25285), *Bifidobacterium adolescentis* (DSMZ 20087), *Bifidobacterium animalis* spp. *lactis* (DSMZ 10140), *Bifidobacterium bifidum* (ATCC 29521), *B. breve* (ATCC 15700), *B. longum* ssp. *longum* (DSMZ 20219), *Lactobacillus casei* (DSMZ 20011), *Lactobacillus reuteri* (CCM 3625), and *Lactobacillus rhamnosus* (CCM 7091). As the maintenance and growth medium, Mueller–Hinton broth (Oxoid, Basingstoke, UK) was used for bacteria that grow aerobically (*E. faecalis* supp. 1% glucose, *V. parahaemolyticus* supp. 3% NaCl). The anaerobic bacteria (clostridia and bifidobacteria), including facultative species (lactobacilli), were cultured in Wilkins–Chalgren broth (Oxoid, Basingstoke, UK) supplemented with 5 g/L soya peptone and 0.5 g/L cysteine.

#### *4.3. Cell Cultures*

One representative normal intestinal cell line (FHs 74 Int (ATCC CCL 241)) and two cancer intestinal cell lines (Caco-2 (ATCC HTB 37) and HT29 (ATCC HTB 38)) were purchased from ATCC (Rockville, MD, USA). Normal cells were cultured in Hybri-Care medium supplemented with 10% fetal bovine serum, 1% sodium bicarbonate, 1% non-essential amino acids, 30 ng/mL of epidermal growth factor, and 1% penicillin-streptomycin solution (10,000 units/mL and 100 mg/mL, respectively). The cancer cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 1% sodium pyruvate, 10% fetal bovine serum, 1% sodium bicarbonate, 1% non-essential amino acids, and 1% penicillin-streptomycin solution (10,000 units/mL and 100 mg/mL, respectively) (all purchased from Sigma-Aldrich, Prague, Czech Republic). The cultures were incubated at 37 ◦C and 5% CO2. The culture medium was replaced every 2–3 days and cells were passaged every 7 days.

#### *4.4. Antibacterial Assay*

The growth-inhibitory activities of the test compounds against aerobic and anaerobic bacterial strains were evaluated by the broth microdilution method using 96-well microtiter plates, following the protocols of CLSI guidelines [48] and Hecht et al. [49], respectively. Prior to testing, the strains were sub-cultured in the appropriate media at 37 ◦C for 24 h. Obligate anaerobes and lactobacilli were cultured for 48 h using Whitley A35 Anaerobic Workstation (Don Whitley Scientific, Bingley, UK). The anaerobic conditions were created by the supply of a standard anaerobic gas mixture of 10% H2, 10% CO2, and 80% N<sup>2</sup> (Linde Gas, Prague, Czech Republic). Test agents were diluted 2-fold in appropriate growth media using the Freedom EVO 100 automated pipetting platform (Tecan, Männedorf, Switzerland) or multichannel pipette (Eppendorf, Hamburg, Germany) (initial concentration of 512 µg/mL). After the bacterial cultures reached an inoculum density of 1.5 × 10<sup>8</sup> CFU/mL by 0.5 McFarland standard using Densi-La-Meter II (Lachema, Brno, Czech Republic), the 96-well plates were inoculated (5 µL/well). The plates containing the volatile compound, 8-hydroxyquinoline, were covered using EVA capmats (Micronic, Lelystad, Netherlands) after inoculation to prevent evaporation [50]. Bacterial cultures in microplates were incubated by employing the same protocols as used for their cultivation prior to the test. The optical density of the cultures was measured at 405 nm (OD450 nm) using a Cytation 3 Imaging Reader (BioTek, Winooski, VT, USA) before and after the growth. The lowest concentration (µg/mL) of test compounds at which the bacterial growth was inhibited by ≥80% was defined as MIC. All tests were performed as three independent experiments each carried out in triplicate. The data are presented as median/mode. As a result of experiments performed without dissolved test compounds, DMSO and 96% ethanol (both from Sigma-Aldrich, Prague, Czech Republic) did not inhibit bacterial growth of any strain at the tested concentrations (≤1%).

#### *4.5. Cytotoxicity Assay*

The antiproliferative activities of test compounds against normal and cancer intestinal lines were evaluated using the modified thiazolyl blue tetrazolium bromide (MTT) cytotoxicity assay developed by Mosmann et al. [51]. The cancer (2.5 × 10<sup>3</sup> ) and normal intestinal (2.5 × 10<sup>5</sup> ) cells were seeded in a 96-well microtiter plate for 24 h. Cells were incubated with two-fold serially diluted test compounds (0.25–512 µg/mL) for 72 h. Plates containing 8-hydroxyquinoline were covered using EVA capmats. Next, the cells were incubated with MTT reagent (1 mg/mL) (Sigma-Aldrich, Prague, Czech Republic) in DMEM or Hybri-Care medium for an additional 2 h at 37 ◦C and 5% CO2. The medium with MTT was removed and the intracellular formazan product was dissolved in 100 µL of DMSO. The absorbance was measured at 555 nm using a Tecan Infinite M200 spectrometer (Tecan, Männedorf, Switzerland), and the percentage of viability was calculated when compared to an untreated control. Antiproliferative activity of the test compounds was represented as IC<sup>50</sup> (µg/mL). Three independent experiments (two replicates each) were performed for every test. Data are presented as mean ± standard deviation. DMSO was used as a positive control at the highest concentration with and without EVA capmats. The solvents did not affect the viability of normal and cancer intestinal cell lines at the tested concentration (≤1%).

#### *4.6. Calculations and Statistics*

For comparison of microbiological and toxicological data, IC<sup>80</sup> was calculated as equivalent to the MIC endpoint, defined as 80% bacterial growth inhibition [52]. Subsequently, *x-*MIC, *x-*IC50, and *x-*IC<sup>80</sup> values (±standard deviations) were calculated to quantify the inhibitory activity of test compounds against diarrheagenic/probiotic bacteria and cancer cells, respectively. After that, SIa (normal intestinal cells/ diarrheagenic strains), SIb (probiotic/diarrheagenic strains), and SIc (normal/cancer intestinal cells) was calculated using the formulas below.

$$\text{SIa} = \log \left( \mathbf{X}\_1 / \mathbf{Y}\_1 \right)\_\prime$$

$$\text{SIb} = \log \left( \mathbf{X}\_2 / \mathbf{Y}\_1 \right)$$

$$\text{SIc} = \log \left( \mathbf{X}\_3 / \mathbf{Y}\_2 \right)$$

where: X<sup>1</sup> = IC<sup>80</sup> against FHs 74 Int; X<sup>2</sup> = *x-*MIC against probiotic strains; X<sup>3</sup> = IC<sup>50</sup> against FHs 74 Int; Y<sup>1</sup> = *x-*MIC against diarrheagenic strains; Y<sup>2</sup> = *x-*IC<sup>50</sup> against cancer intestinal cells. The SI values > 0 and <0 indicate selective toxicity against diarrheagenic strains/cancer cell lines and probiotic strains/normal cell lines, respectively.

The correlation between the combination of activities revealed by test compounds and their chemical classes was analyzed using PCA with Statistica 13 software [53]. All data for particular activities were grouped into four types of targets (cancer cells, diarrheagenic strains, normal cells, and probiotic strains) using MIC and IC<sup>80</sup> values. There was no adjustment of PCA parameters for this analysis. For the calculation of each *x-*MIC, *x-*IC50, *x-*IC<sup>80</sup> and for PCA, values greater than the maximum tested concentration (512 µg/mL) were replaced with 1024 µg/mL.

#### **5. Conclusions**

In summary, ciprofloxacin, 8-hydroxyquinoline alkaloids (chloroxine and nitroxoline), tetracycline, and zinc pyrithione exhibited a significant selective growth-inhibitory activity against diarrheagenic bacteria with lowered toxicity to probiotic bacteria in vitro. 8-Hydroxyquinoline, chloroxine, nitroxoline, sanguinarine, and zinc pyrithione also exhibited a strong cytotoxic effect, whereas the antiproliferative action of 8-hydroxyquinoline and sanguinarine were selective to cancer intestinal cells. These findings indicate that 8-hydroxyquinoline alkaloids and metal-pyridine derivative complexes are chemical structures with promising bioactive properties in terms of in vitro selective antibacterial and anticancer activities which could be utilized in future chemotaxonomic investigation of antidiarrheal medicinal

plants and their bioactive components. These could be further investigated as possible new chemotherapeutic agents against diarrheal infections and associated intestinal cancer diseases. However, in vivo studies on the toxicity of these compounds with more complex animal models will be needed before their consideration to be used for this purpose.

**Author Contributions:** Conceptualization, T.K. and L.K.; methodology, T.K., L.K., and I.D.; software, M.P.; validation, L.K., I.D., H.S., M.P., and E.S.; formal analysis, T.K., L.K., I.D., H.S., M.P., and E.S.; investigation, T.K., L.K., I.D., H.S., and E.S.; resources, L.K., H.S., and E.S.; data curation, T.K., L.K., I.D., and M.P.; writing—original draft preparation, T.K. and L.K.; writing—review and editing, I.D., H.S., M.P., and E.S.; visualization, T.K. and M.P.; supervision, L.K.; project administration, L.K.; funding acquisition, L.K. and E.S. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Czech University of Life Sciences Prague, project IGA 20205001, and European Regional Development Fund, project CZ.02.1.01/0.0/0.0/16\_019/0000845.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **3-Amino-5-(indol-3-yl)methylene-4-oxo-2 thioxothiazolidine Derivatives as Antimicrobial Agents: Synthesis, Computational and Biological Evaluation**


Received: 27 July 2020; Accepted: 28 August 2020; Published: 1 September 2020

**Abstract:** Herein we report the design, synthesis, computational, and experimental evaluation of the antimicrobial activity of fourteen new 3-amino-5-(indol-3-yl) methylene-4-oxo-2-thioxothiazolidine derivatives. The structures were designed, and their antimicrobial activity and toxicity were predicted in silico. All synthesized compounds exhibited antibacterial activity against eight Gram-positive and Gram-negative bacteria. Their activity exceeded those of ampicillin and (for the majority of compounds) streptomycin. The most sensitive bacterium was *S. aureus* (American Type Culture Collection ATCC 6538), while *L. monocytogenes* (NCTC 7973) was the most resistant. The best antibacterial activity was observed for compound **5d** (Z)-N-(5-((1H-indol-3-yl)methylene)-4-oxo-2-thioxothiazolidin-3-yl)-4-hydroxybenzamide (Minimal inhibitory concentration, MIC at 37.9–113.8 µM, and Minimal bactericidal concentration MBC at 57.8–118.3 µM). Three most active compounds **5d, 5g,** and **5k** being evaluated against three resistant strains, Methicillin resistant *Staphilococcus aureus* (MRSA), *P. aeruginosa,* and *E. coli*, were more potent against MRSA than ampicillin (MIC at 248–372 µM, MBC at 372–1240 µM). At the same time, streptomycin (MIC at 43–172 µM, MBC at 86–344 µM) did not show bactericidal activity at all. The compound **5d** was also more active than ampicillin towards resistant *P. aeruginosa* strain. Antifungal activity of all compounds exceeded those of the reference antifungal agents bifonazole (MIC at 480–640 µM, and MFC at 640–800 µM) and ketoconazole (MIC 285–475 µM and MFC 380–950 µM). The best activity was exhibited by compound **5g**. The most sensitive fungal was *T. viride* (IAM 5061), while *A. fumigatus* (human isolate*)* was the most resistant. Low cytotoxicity against HEK-293 human embryonic kidney cell line and reasonable selectivity indices were shown for the most active compounds **5d**, **5g**, **5k**, **7c** using thiazolyl blue tetrazolium bromide MTT assay. The docking studies indicated a probable involvement of *E. coli* Mur B inhibition in the antibacterial action, while CYP51 inhibition is likely responsible for the antifungal activity of the tested compounds.

**Keywords:** indole; thioxothiazolidine; antibacterial activity; antifungal activity; computer-aided prediction; docking; Mur B; CYP 51

#### **1. Introduction**

Infectious diseases affect large populations and cause significant morbidity and mortality [1]. They represent a global indirect load on public health security and an impact on socio-economic stability worldwide. Bacterial, fungal, and viral infections have monopolized the dominant factors of death and disability of millions of humans for centuries. They are presently plaguing and even ravaging populations worldwide each year with performances far surpassing wars [2].

It should be mentioned that several dozen new infections have grown and affected the health of billions of people over the world, mainly in developing countries [3]. Unfortunately, there are no successful pharmaceuticals or vaccines for many of these new infections [3].

The treatment of infectious disease is still an imperative and demanding problem due to the growing number of multi-drug resistant pathogens, especially Gram-positive bacteria. Due to this, the lack of effective antimicrobial drugs, morbidity, and mortality notably increased [4].

Drug resistance causes vast human suffering, and now it is one of the most significant challenges of the twenty-first century. Species such as the methicillin-resistant *S. aureus* and vancomycin-resistant enterococci have emerged due to the irrational or overuse of antimicrobial agents [5].

The pathogens, including *Enterococcus faecium*, *Staphylococcus aureus*, *Klebsiella pneumoniae*, *Acinetobacter baumannii*, *Pseudomonas aeruginosa*, and *Enterobacter spp*. which also called ESCAPE pathogens, are of particular importance since they play a significant role affecting several human organs including the lung and urinary system. Besides, they exhibited increased resistance to clinically used antibiotics [6].

Numerous of these pathogens are Gram-negative bacteria, which are of specific concern due to their resistance of up to 50% against carbapenems that have been reported in some developing countries [6]. Despite the availability of some new antibiotics against Gram-positive pathogens, no treatment of these pathogens with a new class of compounds has been introduced in the last 40 years. Therefore, to overcome the resistance, the discovery of safer and more effective antimicrobial agents with a different mechanism of action is still an urgent need [7].

The interest in thiazolidine-based compounds attracted the attention of medicinal chemists, and a plethora of them have been studied to evaluate pharmacological properties [8–10]. Despite the appearance of some controversial opinions regarding the analysis of the molecular mechanism of their action, prominent representatives among the developed drug-like molecules are thiazolidinone derivatives [11,12] since they are a valuable source of building blocks for the development of novel molecules [13–15].

N-(4-oxo-2-thioxothiazolidin-3-yl)carboxamides exhibit antimicrobial [16–20] and antitumor [21–23] actions, are dual COX-1/2 and 5-LOX inhibitors [24,25], non-nucleoside inhibitors of Hepatitis C NS5b RNA polymerase [26,27] and HIV-1 reverse transcriptase inhibitors [28].

The combination of the thiazolidinone ring with other pharmacologically promising heterocycles has been a warranted approach for developing new "drug-like" molecules with the desired activity profile [29–31]. Our previous studies showed that thiazolidinone core with indole fragment in one molecule gave the compounds with high antimicrobial activity [19].

On the other hand, indole derivatives represent another scaffold widely spread in nature with a broad spectrum of biological activities. The indole ring was found not only in natural compounds but also in diverse semisynthetic and synthetic drug-like molecules [32,33]. They exhibit antimicrobial [34–39], anti-inflammatory [40,41], COX inhibitory [42,43] anticancer [44–46], antiviral [47,48], anti-HIV [49,50], and antidiabetic [51] activities. Among the natural compounds containing the indolene fragment, several imidazoline and imidazolidine alkaloids are known, which have a wide spectrum of biological activity, including antibacterial. Thus, indole-containing azahydantoins 1-6 from sponges and streptomycetes have a potent antibacterial and antiseptic action (Figure 1) [52–54]:

**Figure 1.** Structure of indole-containing azahydantoins 1-5 from sponges and streptomycetes.

It is also known that synthetic thiohydantoin (rhodanine) analogs **7**, **8 (**Figure 2**)**, exhibit pronounced antibacterial properties [55].

**Figure 2.** Synthetic thiohydantoin analogues.

Therefore, the design and development of hybrid molecules combining thiazolidinone and indole cores in the same structure is a promising approach. Taking into account all issues mentioned above and encouraging results obtained in our earlier studies [19], in this paper, we present the synthesis and biological evaluation of new (1H-indole-3-yl-methylene)-4-oxo-2-thioxothiazolidin derivatives with potent antimicrobial activity.

#### **2. Results and Discussion**

#### *2.1. In Silico Antimicrobial Activity Estimation*

#### 2.1.1. Antibacterial Activity

Using AntiBac-Pred [56] one of the predictive web services of Way2Drug platform [57], activity against at least one strain of bacteria was predicted for each of the fourteen designed

compounds with Pa-Pi values in the range from 0.001 to 0.309. According to the prediction results, the highest probability of antibacterial activity against the *Bacillus subtilis subsp. subtilis* str. 168 was estimated for derivatives **7a** and **5b** (Pa-Pi values are 0.309 and 0.305, respectively).

Similarly, we estimated in silico the probability of antibacterial activity for the reference drugs streptomycin and ampicillin. For both reference drugs, wide antibacterial action was predicted. For the top-10 predictions of streptomycin Pa-Pi values vary from 0.905 to 0.947; for ampicillin—from 0.712 to 0.989. Contrary, for relatively new antibacterial agent trifolirhizin, which structure was disclosed only on July 7, 2020 (Clarivate Analytics Integrity [58]), the top-10 predictions Pa-Pi values vary from 0.369 to 0.552.

#### 2.1.2. Antifungal Activity

Using web service AntiFun-Pred [59], activity against at least one of the fungal species was predicted for six of the fourteen studied compounds with Pa-Pi values ranging from 0.001 to 0.112. The results show that among the studied compounds, derivatives 5a (Pa-Pi against *Trichophyton mentagrophytes* equals 0.112) and 7a (Pa-Pi against *Candida equals* 0.101) have better chances to be found active in biological evaluation of the antifungal activity.

The results of in silico antimicrobial activity assessment are given in the supplementary file PASSweb\_results\_13mols.xlsx. Small Pa-Pi values reflect the novelty of the analyzed compounds compared to those included in the PASS training set.

Similarly, for the reference drug ketoconazole wide antifungal action was predicted with Pa-Pi values in the range 0.622–0.812 (top-10 predictions), while for the new antifungal agent drimenin disclosed on 12 June 2020 (Clarivate Analytics Integrity [58]), only two antifungal activity were predicted with Pa-Pi values 0.007 and 0.030.

#### 2.1.3. Acute Rat Toxicity

Using web service based on GUSAR software [60,61], acute rat toxicity with regards to different administration routes was estimated for the studied compounds. LD50 values and toxicity classes are given in Table 1. Most of the predictions indicate that the studied compounds belong to the fifth or fourth rodent toxicity classes.


**Table 1.** In silico assessments of acute rat toxicity.

**Notes:** \*: Calculated for compounds that do not correspond to the model's applicability domain; thus, they are less reliable than unmarked ones. NT: Non-Toxic.

#### *2.2. Chemistry*

The starting N-(4-oxo-2-thioxothiazolidin-3-yl) -carbamides **3a-d** was prepared by reacting the acid hydrazides **1a-d** with trithiocarbonyl diglycolic acid (Scheme 1). The reaction was carried out in a medium of boiling aqueous alcohol. The yield of the products was 83–97%.

**1a, 3a** R = 2-OH, X = Y = CH; **1b, 3b** R = 4-OH, X = Y = CH; **1c, 3c** X = N, R = H, Y = CH; **1d, 3d** Y = N, R = H, X = CH **1a, 3a** R = 2-OH, X = Y = CH; **1b, 3b** R = 4-OH, X = Y = CH; **1c, 3c** X = N, R = H, Y = CH; **1d, 3d** Y = N, R = H, X = CH

**Scheme 1.** Synthesis of initial compounds.

The titled compounds were synthesized according to the process shown in Scheme 2.

#### **Scheme 2.** Synthesis of final compounds.

The reaction of N-(4-oxo-2-thioxothiazolidin-3-yl)carbamides **3a–d** with indole-3-carbaldehydes **4a-d** in acetic acid in the presence of an ammonium acetate catalyst afforded with high yield 5-[(R-1*H*-indol-3-yl)methylene]-4-oxo-2-thioxothiazolidin-3-ylcarbamides **5a–k**, while upon reaction of indole-3-carbaldehydes **4a–d** with 3-morpholino-2-thioxothiazolidin-4-one **6** in the same conditions 5-[(R-1*H*-indol-3-yl) methylene] -3-morpholin-4-yl-2-thioxothiazolidin-4-ones **7a–c** were obtained.

**а а а с**

**а а а с**

All compounds were characterized by IR, <sup>1</sup>H and <sup>13</sup>C NMR spectroscopy. In the IR spectra of compounds **3a–d, 5a–k,** and **7a, 7c**, the carbonyl group of the 4-thiazolidone ring absorbs at 1753.21–1690.53 cm−<sup>1</sup> , and the thiocarbonyl group—at 1608.56–1556.48 cm−<sup>1</sup> . The absorption band of the carbonyl group of the amide fragment of **3a–d** and **5a–k** is located at 1689.56–1654.84 cm−<sup>1</sup> .

In the starting 3-substituted 2-thione-4-thiazolidones, the amide proton NH-CO of the compounds **3a–d** appears as a singlet in the range 11.95-10.91 ppm, and the cyclic methylene group resonates as a singlet or quartet at 4.55–4.48 ppm. etc. In the target products **5a–k**, the amide proton is in the range of 11.85–11.12 ppm. The 5-methylidene proton CH = of compounds **5a–k** and **7a, 7c** resonates in the form of a singlet at 8.20–7.94 ppm, which, according to the literature [9,62], is characteristic of the Z isomer. The singlet NH of the protons of the indole ring appeared in the range 12.31–12.06 ppm.

#### *2.3. Biological Evaluation*

#### 2.3.1. Antibacterial Activity

Compounds **5a–k** and **7a–c** were evaluated for antibacterial activity, by microdilution method to determine the minimal bacteriostatic and bactericidal concentrations. As reference compounds, we used ampicillin and streptomycin, which are both broad-spectrum antibiotics commonly applied to treat different conditions. Antibacterial activity of tested compounds is shown in Table 2 with MIC values in the range of 36.5–211.5 µM and MBC at 73.3–282.0 µM. According to the order of activity which can be presented as: **5d** > **5g** > **5k** > **5j** > **5c** > **5h** > **5e** > **5f** > **5a** > **7c** > **7b** > **5b** > **7a** > **5i** the best activity is achieved for compound **5d** with MIC at 37.9–113.8 µM and MBC at 75.9–151.7 µM. The lowest antibacterial activity was observed for compound **5i** with MIC values in the range of 76.1–152.1 µM and MBC at 152.1–304.2 µM. The most sensitive bacterium appeared to be *S.aureus* (ATCC 6538), *En. cloacae* (ATCC 35,030) was the second most sensitive, while *S.Typhimirium* was the most resistant one. Another resistant strain was Gram-negative bacterium *S. Typhimurium* (ATCC 13,311).

Compound **7b** exhibited good activity against *B. cereus* with MIC and MBC at 41.7 and 83.4 µM respectively. Compound **5d** appeared to be potent against *S. aureus* (ATCC 6538), *P. aeruginosa* (ATCC 27,853), and *En. cloacaei* (ATCC 35,030) with MIC at 37.9 µM and MBC at 75.9 µM. It also showed good activity against *B. cereus* with MIC and MBC at 55.6 and 75.9 µM respectively. Compound **5h** appeared to be potent against *En. cloacae* and *P. aeruginosa* (ATCC 27,853) with MIC and MBC at 39.4 and 78.9 µM. Good activity against these two species and *S.* aureus (ATCC 6538) was also shown by compound 5j (MIC/MBC 58.6/73.1 µM). Good activity against *S. aureus* (ATCC 6538)*,* also exhibited by compound **7b** with MIC at 41.7 µM and MBC at 83.4 µM. On the other hand, compound **5g** exhibited good activity against *En. cloacae* (ATCC 35030)*, S. aureus* (ATCC 6538), and *S. typhimurium* (ATCC 13311) with MIC/MBC values 36.5/73.1 and 53.6/73.1 µM, respectively. It is worth to notice that all compounds appeared to be more potent than ampicillin against all bacteria used and more active than streptomycin against all bacteria except *B. cereus* and *S. typhimurium* (ATCC 13,311).

The structure-activity studies revealed that the most beneficial for antibacterial activity is the presence of hydroxybenzamide (**5d**) on the N-atom of (Z)-5-((5-methoxy-1H-indol-3-yl)methylene)-3 morpholino-2-thioxothiazolidin-4-one. Introduction of the 5-methoxy group to indole ring and replacement of hydroxybenzamide by nicotinamide (**5g**) decreased a little activity while shifting of methoxy group from position 5 to position 6 of indole ring and replacement of nicotinamide by isonicotinamide led to less active compound **5k** compared to compound **5g**.

On the other hand, the isonicotinamide derivative of (Z)-5-((1-methyl-indol-3-yl)methylene)-2 thioxothiazolidin-4-one **(5i)** appeared to be the less active compound. It was observed that for (Z)-5- [(1H-indol-3-yl)methylene]-2-thioxothiazolidin-4-one (**5h**) as well as for (Z)-N-5-[(1-methyl-1Hindol-3-yl)methylene]-4-oxo-2-thioxothiazolidin-4-one (**5g**) derivatives isonicotinamide substituent is endowed with better activity. The opposite was observed for 6-methoxy indole derivatives where more preferable is nicotinamide as a substituent (**5k**). Between methylindole derivatives (**5a, 5f, 5i**), more favorable for activity was nicotinamide substituent (**5f**), followed by benzamide, (**5a**)

while isonicotinamide (**5i**) had a negative effect on antibacterial activity. For -2-hydroxybenzamides derivatives more preferable for antibacterial activity appeared to be 6-methoxy substitution of indole ring (**5c**) followed by methylidole (**5a**) while 5-methoxy substitution on indole ring was negative leading to one of the less active compounds **(5b**). In the case of 3-morpholino-2-thioxothiazolidin-4- one derivatives (**7a–c**), which were among the less active compound, it seems that 5-methoxy substitution on indole ring is preferable than methylindole or indole ring.

Thus, it can be concluded that the most favorable effect on the antibacterial activity of the target compounds is provided by the introduction into the molecule of an unsubstituted indolidene and 6-methoxyindolidene fragment. In addition, the nature of the substituent at position 3 of the thiazolidine ring has a direct influence on the enhancement of the antibacterial action. An increase in the antibacterial effect is observed from the use of 4-hydroxybenzamide and isonicotinamide substitutes.

From all mentioned above, it is evident that the antibacterial activity of these compounds depends not only on substituent and its position in the indole ring but also on substituent on the N-atom of 2-thioxothiazolidin-4-one ring.

**Table 2.** Antibacterial activity of compounds **5a–k** and **7a–c** (MIC/MBC in µM).


MIC–minimal inhibitory concentration, MBC–minimal bactericidal concentration, *B.c.-B.cereus* (clinical isolate), *M.f.-M.flavus* (ATCC 10,240), *S.a.-S.aurues* (ATCC 6538), *l.m.-L.monocytogenes* (NCTC 7973), *E.c.-E.coli* (ATTC 35210, *En.c.-En.cloaca* (ATCC 3503), *P.a.-P.aeruginosa* (ATCC 27,853), *S.T.-S.Typhimurium* (ATCC 13,311).

Three most active compounds were also evaluated against the resistant strains, including MRSA, *P. aeruginosa*, and *E. coli*, (Table 3). From the obtained results, it is evident that all three compounds were more active against MRSA than ampicillin, while streptomycin did not show any bactericidal activity. The compound **5d** was also more active than ampicillin towards resistant *P. aeruginosa* strain.


**Table 3.** Antibacterial activity against resistant strains (MIC/MBC in µM)**.**

#### 2.3.2. Antifungal Activity

All compounds also showed antifungal activity with MIC values ranging from 9.7 to 347.4 µM and MFC at 19.5–694.8 µM.The antifungal activity of compounds is shown in Table 4 and follows the order: **5g** > **7c** > **7b** > **5d** > **5b** > **5e** > **5k** > **5f** > **5j** > **5c** > **5i** > **5a** > **7a** > **5h**. Compound **5g** displayed the best activity with MIC values in the range of 9.7–73.1 µM and MFC at 36.5–146.2 µM, while compound **5h** exhibited the lowest potential with MIC and MFC at 28.9–315.5 µM and 39.4–630.9 µM respectively. It was observed that similar to bacteria, fungi showed different sensitivity towards compounds tested. Thus, the most sensitive fungal strain appeared to be *T. viride* (IAM 5061), while the most resistant filamentous A. fumigatus. The behavior of compounds towards fungi species was different, too.

Several compounds showed very good activity against some species. For example, compound **5d** exhibited good activity against the most resistant *A. fumigatus* (MIC/MFC at .20.2/37.9 µM, while compound **7b** against *T. viride* (IAM 5061), *P. cyclpoium var verucosum* (food isolate) and all Aspergillus species except *A. fumigatus* (human isolate) with MIC at 22.3 µM and MFC at 41.4 µM. Compound **5g** exhibited excellent activity against *T. viride* (IAM 5061) (MIC/MFC at 0.97/1.95 µmol/mL × 10−<sup>2</sup> ). Additionally, good activity was achieved for compound **5g** against *A. versicolor* (ATCC 11730), *A. ochraceus* (ATCC 12066), *P. funiculosum* (ATCC 36839) with MIC and MFC at 19.5 µM and 36.5 µM respectively. Compound **5c** appeared to be potent against *A. ochraceus* (ATCC 12066) and *T. viride* (IAM 5061) (MIC/MFC at 18.8/35.3 µM whereas compound **7c** exhibited very good activity against *T. viride* (IAM 5061)with MIC at 10.7 µM and MFC at 21.3 µM and also good activity against *A. ochraceus* (ATCC 12066) and *P. funiculosum* (ATCC 36839 (MC/MFC at 23.1/39.9 µM. The potential of ketoconazole was at MIC 285-475 µM and MFC at 380–950 µM. Bifonazole displayed MIC at 480-640 µM and MFC at 640–800 µM. It should be mentioned that all compounds appeared to be more potent than ketoconazole and bifonazole. Only compound **7a** against *A. fumigatus (human isolate)* was less active than bifonazole.

According to the analysis of the structure-activity relationships, the most beneficial for antifungal activity is the presence of the 5-methoxy group in indole ring as well as nicotinamide as a substituent of the side chain (**5g**). In contrast, the presence of isonicotinamide in methylindole (**5i)** derivative appeared to be detrimental. Shifting of 5-OMe of compound **5g** to position 6 of indole and replacement of nicotinamide by 2-hydroxybenzamide resulted in compound **5c** with decreased activity. Removal of methoxy group and introduction of morpholino moiety to the N atom of thioxothiazolidinone **(7a)** decreased more activity.

In indole derivatives (**5d**, **5e**, **5h**), the presence of 4-hydroxybenzamide was favorable for antifungal activity, while isonicotinamide substituent had a negative effect. On the contrary, for methylindole derivatives (**5a**, **5f**, **5i**), the negative impact was observed with the presence of 2-hydroxybenzamide, while in the case of the 5-methoxy indole derivatives (**5b**, **5j**) it was the opposite. Finally, for the derivatives with morpholino moiety, the best activity was observed with the presence of the 5-methoxy group in the indole ring. The indole derivative was one of the less potent.

Thus, as in the case of antibacterial activity, antifungal activity depends not only on substitution in the indole ring but also on substituent on the N-atom of the 2-thioxothiazolidinone ring. In the series of (Z)-5-((5-methoxy-1H-indol-3-yl)methylene)-3-morpholino-2-thioxothiazolidin-4-one derivatives the most important structural features which enhanced the antifungal activity are again 4-hydroxybenzamide and 1H-indole moiety as well as nicotinamide and 5- and 6-methoxyindole moieties. On the other hand, in the series of indole 3-methylene morpholino-2-thioxothiazolidin-4-one derivatives, the presence of the 5-OCH3 group in the indole ring enhance the antifungal activity.


**Table 4.** Antifungal activity of compounds **5a–k** and **7a–c** (MIC/MFC in µM).

MIC–minimal inhibitory concentration, MFC–minimal fungicidal concentration*. A.fum.-A.fumigatus* (human isolate), *A.v.-A.versicolor* (ATCC 11730), *A.o.-A.ochraceus* (ATCC 12066), *A.n.-A.niger* (ATCC 6275), *T.v.-T.viride* (IAM 5061), *P.f.-P.funiculosum* (ATCC 36839), *P.o.-P.ochrochloron* (ATCC 9112), *P.v.c.-P.cyclpoium var. verucosum* (food isolate).

#### 2.3.3. Cytotoxicity Assessment

Low toxicity and selectivity of action of antimicrobial compounds is a crucial pre-requisite for further development. Thus, we studied the cytotoxicity of the most active compounds. MTT analysis was performed on the HEK-293 human embryonic kidney cell line. The cells were cultured in DMEM medium supplemented with 10% fetal bovine serum. The cells were inoculated into a 96-well plate at a concentration of 5.10<sup>4</sup> /mL (5.10<sup>3</sup> per well, 100 µL each). After one day of culture, compound preparations were added, and the results were obtained after a 72 h culture period. The compounds were added at four concentrations (25, 50, 100, and 250 µM). Since compound solutions contained DMSO, control cultures containing only DMSO at the final concentration obtained when the appropriate volume of compound solution was added were performed.

Although the compounds do not exhibit statistically significant concentration-dependent toxicity up to 100 µM (Figure 3), they show some toxicity at higher concentrations. The average CC<sup>50</sup> values obtained from three different experiments are given in Tables 5 and 6. The SI index is also shown in Tables 5 and 6.

Compound **5g** and **7c** exhibited the best SI index for anti-fungal activity while compound **5d** exhibited the best SI index for anti-bacterial activity.

We compared the CC<sup>50</sup> values of compounds **5d**, **5k**, **5g**, **7c** with cytotoxicity of the reference drugs obtained in the HEK-293 human embryonic kidney cell line. For antibacterials streptomycin, ampicillin and antifungal bifonazole CC<sup>50</sup> exceeded 100 µM [63,64]; for antifungal ketoconazole CC<sup>50</sup> = 60 µM [65]. Thus, cytotoxicity of the most active compounds in our study is comparable or lower than cytotoxicity of the reference antimicrobial drugs.

**Table 5.** Antibacterial activity (MIC), cytotoxicity (CC50), and selectivity indices (SI) of compounds **5d, 5g, 5k, 7c. μΜ**


**Table 6.** Antifungal activity (MIC), cytotoxicity (CC50), and selectivity indices (SI) of compounds **5d, 5g, 5k, 7c.**


\* 24 h.

μΜ **Figure 3.** MTT assay results for compounds **5d, 5k, 5g, 7c.** According to the results, all compounds did not show statistically significant, concentration-dependent cytotoxicity at concentrations up to 100 µM. The stable decrease in viability observed can be attributed to dimethyl soulfoxide (DMSO,) present at stable concentration at all compound samples.

320

#### *2.4. Docking Studies*

Since the mechanism of antimicrobial action of our compounds is not known, to choose the proteins as potential targets, we based on the literature. It was found that benzothiazole derivatives are mentioned as Gyrase inhibitors [66–68]. On the other hand, according to the literature, thiazolidinones act as MurB inhibitors [69–72]. Furthermore, prediction of the mechanism of action by computer program PASS indicated Thymidylate kinase as the probable antibacterial target. On the other hand, several publications mentioned thiazolidinone and indole derivatives as 14α-lanosterol demethylase inhibitors [73–75]. Thus, taking all these into account, we proposed *E. coli* DNA Gyrase, Thymidylate kinase, and *E. coli* MurB enzymes as antibacterial targets, with CYP51 as the antifungal target.

#### 2.4.1. Docking to Antibacterial Targets

The docking studies revealed that estimated binding energy to *E. coli* DNA Gyrase (−2.59 to −6.54 kcal/mol) as well as to thymidylate kinase (−1.55 to −4.12 kcal/mol), were higher than that to *E. coli* MurB (−7.07 to −10.93 kcal/mol). Therefore, it may be resolved that *E. coli* MurB is the most suitable enzyme where binding scores were consistent with biological activity (Table 7).


**Table 7.** Molecular docking binding energies.

The docking pose of the most active compound **5d** in *E. coli* MurB enzyme showed two favorable hydrogen bond interactions. The first one is between the oxygen atom of the C=O group of the compound and the hydrogen of the side chain of Ser228. The second one between the oxygen atom of -OH group of the compound and the side chain of Arg326 (distances 2.17 Å and 1.99 Å, respectively). The fused rings interact hydrophobically with the residues Tyr189, Asn232, Leu289, Ala123, Leu217, and Arg213, while the benzene ring interacts hydrophobically with the residues Asn50, Ser115, Ile118, Ile121, Gln119 and Glu324 (Figure 4). These interactions stabilize the complex compound-enzyme and play a crucial role in the increased inhibitory activity of compound **5d** Moreover, the hydrogen bond formation with the residue Ser228 is essential for the inhibitory action of the compounds; thus, this residue takes part in the proton transfer at the second stage of peptidoglycan synthesis [76].

The second most active compound, **5g,** also forms the hydrogen bond interaction with the residue Ser228 that explains its high inhibitory action (Figure 2). Detailed analysis of the docking pose of the two most active compounds showed that they similarly bind MurB, and they insert deeper to the binding center of the enzyme than FAD, forming a hydrogen bond with the residue Ser228 (Figure 5).

**Figure 4.** Docked conformation of the most active compound **5d** in *E.coli* MurB **(Left)**. 2D diagrams of the most active compounds **5d** (up) and **5g** (down) in *E.coli* MurB **(Right)**.

The same behavior was observed in the case of docking of the most active compound among 5-(1*H*-indol-3-ylmethylene)-4-oxo-2-thioxothiazolidin-3-yl)alkane carboxylic acids [19] and 5-adamantane thiadiazole-based thiazolidinones [70]. Again, the formation of the hydrogen bond between the C=O group and Ser228 was observed. Thus, the obtained results support previous data [69–72] that MurB maybe is the most appropriate target for the antibacterial activity for this chemical series.

**Figure 5.** Docked conformation of compounds **5d** (green), **5g** (red) and FAD (blue) in *E.Coli* MurB.

α α 2.4.2. Docking to Lanosterol 14α-demethylase of *C. albicans*

α α All the synthesized compounds and the reference drug ketoconazole were docked to lanosterol 14α-demethylase of *C. albicans* (Table 8).



Docking results showed that all the synthesized compounds might bind to CYP51Ca close to those of the reference drug ketoconazole. Compound **5g** is located inside the enzyme alongside to heme group, interacting with the Fe of the heme group of CYP51Ca throughout its atom N of the pyridine ring. Moreover, compound **5g** forms a hydrogen bond between the oxygen of –OCH<sup>3</sup> substituent and the hydrogen of the side chain of Tyr132. Hydrophobic interactions were detected between residues Thr122, Phe126, Tyr132, and Ile131 and the fused rings of the compound **5g**, also between Leu376, Thr311 and the benzene ring of the compound. Furthermore, compound **5g** interacts hydrophobically throughout its benzene ring with the heme group of the enzyme, and also it forms a positive ionizable bond with it (Figure 6). Interaction with the heme group was also observed with the benzene ring of ketoconazole, which forms positive ionizable interactions (Figures 6 and 7). However, compound **5g** forms a more stable complex of the ligand with enzyme indicating its interaction with the Fe, which is probably why compound **5g** showed high antifungal activity.

**Figure 6.** Docked conformation of ketoconazole in lanosterol 14alpha-demethylase of *C*. *albicans* (CYP51ca).

μ

**Figure 7.** Docked conformation of compound **5g** in lanosterol 14alpha-demethylase of *C*. *albicans* (CYP51ca).

It should be mentioned that the tested compounds interact more strongly with the heme group of the enzyme CYP51Ca because the heme's Fe is involved in this interaction. In the case of our previous work [19], the most active compound interacts with the heme but throughout its benzene ring and the –NO<sup>2</sup> group, forming pi and negative ionizable interactions with the heme group, respectively. In the case of 5-adamantane thiadiazole-based thiazolidinones **[72]**, again, the most active compound form positive interactions between the heme group and heterocyclic rings of the compound. Thus, it can be concluded that thiazolidinone derivatives, in general, can interact with the heme of CYP51Ca in the same way as ketoconazole interacts.

#### **3. Materials and Methods**

μ All starting materials were purchased from Merck and used without purification. NMR spectra were determined with Varian Mercury VX-400" (Varian Co., Palo Alto, CA, USA) and AM-300 Bruker 300 MHz. spectrometers in DMSO-d6. MS (ESI) spectra were recorded on an LC-MS system-HPLC Agilent 1100 (Agilent Technologies Inc., Santa, Clara, CA USA) equipped with a diode array detector Agilent LC\MSD SL. Parameters of analysis: Zorbax SB-C18 column (1.8 µM, 4.6–15 mm, PN 821975-932), solvent water–acetonitrile mixture (95:5), 0.1% of aqueous trifluoroacetic acid; eluent flow 3 mL min–1; injection volume 1 µL; IR spectra were recorded on a Vertex 70 Bruker" (Bruker, Karlsruhe, Germany) spectrometer in KBr pellets. Melting points were determined in open capillary tubes and are uncorrected.

#### *3.1. In Silico Biological Activity Evaluation*

Antimicrobial activity and toxicity of the designed compounds have been estimated in silico using web services available on the Way2Drug portal [56]. These services are based on the PASS (Prediction of Activity Spectra for Substances) and GUSAR (General Unrestricted Structure-Activity Relationships) software, which is described in detail elsewhere [60,61]. It is essential to mention that PASS-based services provide the assessments of the compound's activity as the difference between the probabilities for the chemical compound with a particular structure to display activity (Pa) and do not display this activity (Pi). By default, in PASS, all activities with Pa > Pi are considered as probable. High Pa-Pi values reflect the high structural similarity of the analyzed compound to the structures included in the training set with those activities. Since our goal was not finding close analogs of the

earlier discovered antimicrobial agents, we considered compounds with small Pa-Pi values as the promising hits for experimental testing. If the experiment will confirm their activity, there is a chance to find a New Chemical Entity. GUSAR-based service [60,61] provides the quantitative assessment of acute rat toxicity expressed as LD<sup>50</sup> values for four routes of administration: intraperitoneal (IP), intravenous (IV), oral, and subcutaneous (SC).

#### *3.2. Chemistry*

#### 3.2.1. General Procedure for the Preparation of N-(4-oxo-2-thioxothiazolidin-3-yl) carbamides **3a**–**d**

In a round-bottom flask equipped with a reflux condenser, 0.05 mol of trithiocarbonyl diglycolic acid, 0.05 mol of the corresponding hydrazide and alcohol-water mixture (1:1) were placed and boiled for 3 h. The reaction mixture is cooled, the precipitate is filtered off and recrystallized.

**2-Hydroxy-N-(4-oxo-2-thioxothiazolidin-3-yl)benzamide 3a.** Yield 97%; m.p. 104–106 ◦C (CH3COOH-H2O 2:1). IR (cm–1): 3342.48 (OH), 1751.28 (C=O), 1657.74 (C=O), 1608.56 (C=S).). <sup>1</sup>H NMR (400 MHz, DMSO-d6, ppm) δ 11.62–10.91 (br.s, 2H, NH, OH), 7.88 (dd, J = 8.0, 1.6 Hz, 1H, H<sup>6</sup> benzene), 7.52–7.46 (m, 1H, H<sup>3</sup> benzene), 7.05–6.95 (m, 1H, 2H, H<sup>4</sup> +H5, aromatic), 4.48 (q, J = 18.7 Hz, 2H, CH2). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 199.90, 170.25, 164.91, 157.93, 134.62, 129.83, 119.44, 117.19, 115.07, 33.38. Anal. Calcd. for C10H8N2O3S<sup>2</sup> (%): C, 44.77; H, 3.01; N, 10.44; S, 23.90 Found (%):C, 44.88; H, 3.09; N, 10.37; S, 23.95.

**4-Hydroxy-***N***-(4-oxo-2-thioxothiazolidin-3-yl)benzamide 3b.** Yield 87%; m.p. 207–209 ◦C (CH3COOH).). IR (cm–1): 3259.54 (OH), 3166(NH), 1739.71 (C=O), 1667.38(C=O), 1583.48 (C=S). <sup>1</sup>H NMR (400 MHz, DMSO-d6, ppm) δ 11.25 (s, 1H, NH), 10.29 (s, 1H, OH), 7.81 (dd, J = 9.1, 2.3 Hz, 2H, H<sup>2</sup> +H6, benzene), 6.88 (dd, J = 9.1, 2.3 Hz, 2H, H<sup>3</sup> +H5, benzene), 4.51 (s, 2H, CH2). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 200.33, 170.54, 163.93, 161.44, 129.96, 121.44, 115.24, 33.32. Anal. Calcd. for C10H8N2O3S<sup>2</sup> (%): C, 44.77; H, 3.01; N, 10.44; S, 23.90 Found (%):C, 44.69; H, 2.95; N, 10.36; S, 23.81.

**N-(4-Oxo-2-thioxothiazolidin-3-yl)nicotinamide 3c.** Yield 83%; m.p. 190 ◦C decomp.(C2H5OH). IR (cm–1): 1753.21 (C=O), 1687.63 (C=O), 1556.48 (C=S). <sup>1</sup>H NMR (400 MHz, DMSO-d6, ppm) δ 11.95 (s, 1H, NH), 8.97–8.73 (m, 2H, H<sup>2</sup> +H4, pyridine), 7.99–7.70 (m, 2H, H<sup>5</sup> +H6, pyridine), 4.55 (s, 2H, CH2).). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 199.81, 170.20, 163.25, 150.76, 137.93, 121.31, 119.56, 33.55. Anal. Calcd. for C9H7N3O2S<sup>2</sup> (%): C, 42.68; H, 2.79; N, 16.59; S, 25.32 Found (%):C, 42.79; H, 2.70; N, 16.48; S, 25.45.

**N-(4-Oxo-2-thioxothiazolidin-3-yl)isonicotinamide 3d**. Yield 85%; m.p. 193 ◦C decomp. (C2H5OH). IR (cm–1): 1753.21(C=O), 1678.95 (C=O), 1556.48 (C=S).1H NMR (400 MHz, DMSO-d6, ppm) δ 11.95 (s, 1H, NH), 8.87–8.79 (m, 2H, H<sup>3</sup> +H5, pyridine), 7.85–7.80 (m, 2H, H<sup>2</sup> +H6, pyridine), 4.55 (s, 2H, CH2). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 199.81, 170.19, 163.25, 150.76, 137.93, 121.39, 119.56, 33.58. Anal. Calcd. for C9H7N3O2S<sup>2</sup> (%): C, 42.68; H, 2.79; N, 16.59; S, 25.32 Found (%):C, 42.77; H, 2.85; N, 16.76; S, 25.26.

3.2.2. General Procedure 5-[(R-1H-indol-3-ylmethylene)-4-oxo-2-thioxothiazolidin-3-yl] carbamides **5a–k** and 5-(R-1H-indol-3-ylmethylene)-3-morpholin-4-yl-2-thioxothiazolidin-4-ones **7a–c**

In a round-bottom flask equipped with a reflux condenser, 2.5 mmol of 3-substituted 2-thioxo-4-oxothiazolidine **3a-d** or **6**, 3.3 mmol of the corresponding aldehyde **1a-d,** 2.5 mmol of ammonium acetate and 5 mL of acetic acid are placed. The reaction mixture is boiled for 2 h, cooled, the precipitate is filtered off, washed with acetic acid and water, dried and recrystallized.

### **2-Hydroxy-***N***-{(5***Z***)-5-[(1-methyl-1***H***-indol-3-yl)methylene]-4-oxo-2-thioxothiazolidin-3-**

**yl}benzamide 5a.** Yield 98%; m.p. 265–267 ◦C (DMFA-CH3COOH). IR (cm–1): 3272.08 (OH), 1700.17 (C=O), 1656.77 (C=O), 1588.3 (C=C), 1573.84 (C=S). <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) δ 11.40 (s, 1H, NH-CO), 11.31 (s, 1H, OH), 8.12 (s, 1H, CH=), 8.01 (d, *J* = 7.9 Hz, 1H, H<sup>6</sup> benzene), 7.91 (d, *J* = 4.2 Hz, 2H, H<sup>4</sup> +H7, indole), 7.54–7.41 (m, 2H, H<sup>3</sup> benzene + H<sup>2</sup> indole), 7.37–7.22 (m, 2H, H<sup>5</sup> +H6, indole), 6.97 (dd, *J* = 17.3, 8.1 Hz, 2H, H<sup>4</sup> +H5, benzene), 4.00 (s, 3H, CH3N). <sup>13</sup>C NMR

(101 MHz, DMSO, ppm) δ 189.71, 164.99, 163.08, 158.00, 136.96, 134.65, 134.45, 129.90, 127.29, 127.01, 123.54, 122.01, 119.46, 118.78, 117.22, 115.13, 111.01, 109.93, 33.50. ESI-MS [m/z]: [M + H]<sup>+</sup> = 411.0; [M − H]<sup>−</sup> = 408.2. Anal. Calcd. for C20H15N3O3S<sup>2</sup> (%): C, 58.66; H, 3.69; N, 10.26; S, 15.66 Found (%): 58.75 H, 3.62; N, 10.21; S, 15.52.

**2-Hydroxy-***N***-{(5***Z***)-5-[(5-methoxy-1***H***-indol-3-yl)methylene]-4-oxo-2-thioxothiazolidin-3 yl}benzamide 5b.** Yield 77%; m.p. 239–241 ◦C (DMFA-CH3COOH).). IR (cm–1): 3234.47 (OH), 1699.21 (C=O), 1654.84 (C=O), 1585.41 (C=C, C=S). <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) δ 12.15 (s, 1H, NH), 11.45 (s, 1H, NH-CO), 11.35 (s, 1H, OH), 8.18 (s, 1H, CH=), 8.00 (d, *J* = 7.8 Hz, 1H, H<sup>6</sup> benzene), 7.74 (d, *J* = 2.2 Hz, 1H, H<sup>4</sup> indole), 7.50–7.33 (m, 3H, H<sup>3</sup> benzene + H<sup>2</sup> +H7, indole), 7.04–6.90 (m, 2H, H<sup>4</sup> +H5, benzene), 6.83 (d, *J* = 8.4 Hz, 1H, H<sup>6</sup> indole), 3.87 (s, 3H, CH3O <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 189.72, 165.10, 163.07, 158.10, 155.45, 134.64, 131.12, 131.09, 129.80, 128.22, 127.84, 119.43, 117.25, 115.07, 113.67, 113.38, 111.09, 110.44, 100.58, 55.85. ESI-MS [m/z]: [M + H]<sup>+</sup> = 426.0; [M − H]<sup>−</sup> = 424.0. Anal. Calcd. for C20H15N3O4S<sup>2</sup> (%): C, 56.46; H, 3.55; N, 9.88; S, 15.07 Found (%):C, 56.57; H, 3.49; N, 9.96; S, 15.15.

**2-Hydroxy-***N***-{(5***Z***)-5-[(6-methoxy-1***H***-indol-3-yl)methylene]-4-oxo-2-thioxothiazolidin-3 yl}benzamide 5c.** Yield 80%; m.p. 268–270 ◦C (DMFA-CH3COOH). IR (cm–1): 3227.72 (OH), 1705.96 (C=O), 1670.27 (C=O), 1591.2 (C=C), 1576.73 (C=S). <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) δ 12.06 (s, 1H, NH), 11.45 (s, 1H, NH-CO),), 11.34 (s, 1H, OH), 8.10 (s, 1H, CH=), 7.99 (d, *J* = 7.9 Hz, 1H, H<sup>6</sup> benzene), 7.75 (d, *J* = 8.7 Hz, 1H, H<sup>4</sup> indole), 7.69 (d, *J* = 1.9 Hz, 1H, H<sup>3</sup> benzene), 7.48-7,42 (m, 1H, H<sup>2</sup> indole), 7.05–6.91 (m, 3H, H<sup>7</sup> indole +H<sup>4</sup> + H5, benzene), 6. 83 (d, *J* = 8.5 Hz, 1H, H<sup>5</sup> indole), 3.84 (s, 3H, CH3O13C NMR (101 MHz, DMSO, ppm) δ 189.71, 165.00, 163.10, 158.01, 156.93, 137.32, 134.64, 130.31, 129.87, 127.93, 120.68, 119.49, 119.44, 117.22, 115.11, 111.63, 111.13, 111.06, 95.41, 55.28. ESI-MS [m/z]: [M + H]<sup>+</sup> = 426.0; [M − H]<sup>−</sup> = 424.0.. Anal. Calcd. for C20H15N3O4S<sup>2</sup> (%): C, 56.46; H, 3.55; N, 9.88; S, 15.07 Found (%):C, 56.39; H, 3.51; N, 9.80; S, 15.01.

**4-Hydroxy-***N***-[(5***Z***)-5-(1***H***-indol-3-ylmethylene)-4-oxo-2-thioxothiazolidin-3-yl]benzamide 5d.** Yield 90%; m.p. > 275 ◦C (DMFA-CH3COOH). IR (cm–1): 3369.48 (OH), 3225.79 (NH), 1694.38 (C=O), 1668.35 (C=O), 1591.2 (C=C), 1573.84 (C=S). <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) δ 12.23 (s, 1H, NH), 11.12 (s, 1H, NH-CO), 9.89 (s, 1H, OH), 8.13 (s, 1H, CH=), 7.92–7.81 (m, 3H, H<sup>2</sup> + H6, benzene + H<sup>4</sup> indole), 7.79 (d, *J* = 3.0 Hz, 1H, H<sup>7</sup> indole), 7.53–7.45 (m, 1H, H<sup>2</sup> indole), 7.28–7.15 (m, 2H, H<sup>5</sup> + H6, indole), 6.85 (d, *J* = 8.7 Hz, 2H, H<sup>3</sup> +H5, benzene). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 190.12, 164.06, 163.36, 161.49, 136.48, 131.20, 129.97, 127.80, 126.77, 123.49, 121.69, 121.50, 118.64, 115.30, 112.61, 111.16, 110.97. ESI-MS [m/z]: [M + H]<sup>+</sup> = 396.0; [M − H]<sup>−</sup> = 394.0. Anal. Calcd. for C19H13N3O3S<sup>2</sup> (%): C, 57.71; H, 3.31; N, 10.63; S, 16.22 Found (%):C, 57.62; H, 3.37; N, 10.55; S, 16.16.

*N***-[(5***Z***)-5-(1***H***-Indol-3-ylmethylene)-4-oxo-2-thioxothiazolidin-3-yl]nicotinamide 5e.** Yield 90%; m.p. > 275 ◦C (DMFA-CH3COOH). IR (cm–1): 3485.2 (NH), 1696.31 (C=O), 1674.13 (C=O), 1596.98 (C=C), 1577.7 (C=S). <sup>1</sup>H-NMR (500 MHz, DMSO-d6, ppm) δ 12.31 (s, 1H, NH), 11.75 (s, 1H, NH-CO), 9.15 (d, *J* = 1.7 Hz, 1H, H<sup>4</sup> pyridine), 8.77 (dd, *J* = 4.8, 1.4 Hz, 1H, H<sup>2</sup> pyridine), 8.33 (d, *J* = 8.0 Hz, 1H, H<sup>6</sup> pyridine), 8.17 (s, 1H, CH=), 7.90 (d, *J* = 7.6 Hz, 1H, H<sup>5</sup> pyridine), 7.84 (d, *J* = 3.0 Hz, 1H, H<sup>4</sup> indole), 7.57–7.48 (m, 2H, H<sup>2</sup> +H7, indole), 7.27–7.18 (m, 2H, H<sup>5</sup> +H6, indole). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 189.54, 163.32, 162.94, 150.78, 137.89, 136.41, 131.49, 128.38, 126.78, 123.56, 121.78, 121.38, 118.68, 112.65, 110.98, 110.72. ESI-MS [m/z]: [M + H]<sup>+</sup> = 381.0; Anal. Calcd. for C18H12N4O2S<sup>2</sup> (%): C, 56.83; H, 3.18; N, 14.73; S, 16.86 Found (%):C, 56.71; H, 3.24; N, 14.80; S, 16.79.

*N***-{(5***Z***)-5-[(1-Methyl-1***H***-indol-3-yl)methylene]-4-oxo-2-thioxothiazolidin-3-yl}nicotinamide 5f.** Yield 94%; m.p. 270–272 ◦C (DMFA-CH3COOH). IR (cm–1): 3241.22 (NH), 1706.92 (C=O), 1681.85 (C=O), 1589.27 (C=C), 1572.87 (C=S). <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) δ 11.75 (s, 1H, NH-CO), 9.16 (d, *J* = 1.9 Hz, 1H, <sup>H</sup><sup>4</sup> pyridine), 8.77 (dd, *J* = 4.8, 1.3 Hz, 1H, H<sup>2</sup> pyridine), 8.36–8.31 (m, 1H, H<sup>6</sup> pyridine), 8.12 (s, 1H, CH=), 7.96 (s, 1H, H<sup>5</sup> pyridine), 7.91 (d, 1H, *J* = 7.9 Hz, H<sup>4</sup> indole), 7.58–7.47 (m, 2H, H<sup>2</sup> + H7, indole), 7.32 (t, *J* = 7.5 Hz, 1H, H<sup>6</sup> indole), 7.26 (t, *J* = 7.4 Hz, 1H, H<sup>5</sup> indole), 4.00 (s, 3H, CH3N). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 189.64, 163.34, 163.01, 153.40, 148.60, 136.99, 135.59, 134.68, 127.57, 127.30, 126.77, 123.97, 123.60, 122.10, 118.80, 111.08, 110.55, 109.95, 33.54. ESI-MS [m/z]: [M + H]<sup>+</sup> = 395.0; [M − H]<sup>−</sup> = 394.0. Anal. Calcd. for C19H14N4O2S<sup>2</sup> (%): C, 57.85; H, 3.58; N, 14.20; S, 16.26 Found (%):C, 57.94; H, 3.51; N, 14.15; S, 16.35.

*N***-{(5***Z***)-5-[(5-Methoxy-1***H***-indol-3-yl)methylene]-4-oxo-2-thioxothiazolidin-3-yl}nicotinamide 5g.** Yield 00%; m.p. 199–201 ◦C. IR (cm–1): 3254.72 (NH), 1718.49 (C=O), 1681.85 (C=O), 1585.41 (C=C), 1576.73 (C=S). <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) δ 12.17 (s, 1H, NH), 11.72 (s, 1H, NH-CO), 9.17 (s, 1H, H<sup>4</sup> pyridine), 8.77 (d, *J* = 3.0 Hz, 1H, H<sup>2</sup> pyridine), 8.35 (d, *J* = 7.5 Hz, 1H, H<sup>6</sup> pyridine), 8.19 (s, 1H, CH=), 7.74 (s, 1H, H<sup>5</sup> pyridine), 7.59–7.49 (m, 1H, H<sup>2</sup> indole), 7.46–7.30 (m, 2H, H<sup>4</sup> +H<sup>7</sup> indole), 6.83 (d, *J* = 8.5 Hz, 1H, H<sup>6</sup> indole), 3.86 (s, 3H, CH3O). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 189.67, 163.30, 163.01, 155.51, 153.36, 148.61, 135.57, 131.33, 131.15, 128.76, 127.86, 126.82, 123.94, 113.69, 113.41, 111.12, 110.01, 100.64, 55.54. ESI-MS [m/z]: [M + H]<sup>+</sup> = 411.0; [M − H]<sup>−</sup> = 409.0. Anal. Calcd. for C19H14N4O3S<sup>2</sup> (%): C, 55.60; H, 3.44; N, 13.65; S, 15.62 Found (%): C, 55.49; H, 3.39; N, 13.58; S, 15.67.

*N***-[(5***Z***)-5-(1***H***-Indol-3-ylmethylene)-4-oxo-2-thioxothiazolidin-3-yl]isonicotinamide 5h.** Yield 86%; m.p. > 275 ºC Yield 86%; m.p. > 275 ◦C (DMFA-CH3COOH). IR (cm–1): 3196.86 (NH), 1718.49 (C=O), 1672.2 (C=O), 1594.09 (C=C), 1576.73 (C=S). <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) δ 12.27 (s, 1H, NH), 11.79 (s, 1H, NH-CO), 8.78 (d, *J* = 5.8 Hz, 2H, H<sup>2</sup> +H6, pyridine), 8.17 (s, 1H, CH=), 7.94–7.86 (m, 3H, H<sup>3</sup> +H5, pyridine +H<sup>4</sup> indole), 7.82 (s, 1H, H<sup>7</sup> indole), 7.51 (d, *J* = 7.1 Hz, 1H, H<sup>2</sup> indole), 7.30–7.15 (m, 2H, H<sup>5</sup> +H6, indole). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 189.53, 163.32, 162.94, 150.76, 137.94, 136.42, 131.46, 128.34, 126.77, 123.54, 121.76, 121.38, 118.66, 112.65, 110.99, 110.78. ESI-MS [m/z]: [M + H]<sup>+</sup> = 381.0; [M − H]<sup>−</sup> = 379.0. Anal. Calcd. for C18H12N4O2S<sup>2</sup> (%): C, 56.83; H, 3.18; N, 14.73; S, 16.86 Found (%):C, 56.89; H, 3.26; N, 14.65; S, 16.88.

*N***-{(5***Z***)-5-[(1-Methyl-1***H***-indol-3-yl)methylene]-4-oxo-2-thioxothiazolidin-3 yl}isonicotinamide 5i.** Yield 95%; m.p. 269–271 ◦C (DMFA-CH3COOH). IR (cm–1): 3217.11 (NH), 1710.78 (C=O), 1674.13 (C=O), 1587.34 (C=C), 1570.95 (C=S).1H-NMR (300 MHz, DMSO-d6, ppm) δ 11.81 (s, 1H, NH-CO), 8.79 (d, *J* = 5.9 Hz, 2H, H<sup>2</sup> +H6, pyridine), 8.13 (s, 1H, CH=), 7.99–7.86 (m, 4H, H<sup>3</sup> +H5, pyridine + H<sup>4</sup> +H7, indole), 7.51 (d, *J* = 7.9 Hz, 1H, H<sup>2</sup> indole), 7.37–7.20 (m, 2H, H<sup>5</sup> +H6, indole), 4.00 (s, 3H, CH3N). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 189.49, 163.32, 162.90, 150.78, 137.88, 137.00, 134.73, 127.68, 127.30, 123.61, 122.12, 121.38, 118.81, 111.09, 110.46, 109.95, 33.55. ESI-MS [m/z]: [M + H]<sup>+</sup> = 395.0; [M − H]<sup>−</sup> = 393.0. Anal. Calcd. for C19H14N4O2S<sup>2</sup> (%): C, 57.85; H, 3.58; N,

### 14.20; S, 16.26 Found (%):C, 57.78; H, 3.53; N, 14.28; S, 16.17.

*N***-{(5***Z***)-5-[(5-Methoxy-1***H***-indol-3-yl)methylene]-4-oxo-2-thioxothiazolidin-3 yl}isonicotinamide 5j.** Yield 89%; m.p. 261–263 ºC (CH3COOH).). IR (cm–1): 3199.75 (NH), 1706.92 (C=O), 1676.06 (C=O), 1588.3 (C=C, C=S). <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) δ 12.20 (s, 1H, NH), 11.83 (s, 1H, NH-CO), 8.78 (d, *J* = 5.4 Hz, 2H, H<sup>2</sup> +H6, pyridine), 8.20 (s, 1H, CH=), 7.90 (d, *J* = 5.4 Hz, 2H, H<sup>3</sup> +H5, pyridine), 7.76 (d, *J* = 3.0 Hz, 1H, H<sup>4</sup> indole), 7.44–7.33 (m, 2H, H<sup>2</sup> +H7, indole), 6.83 (dd, *J* = 8.9, 1.7 Hz, 1H, H<sup>6</sup> indole), 3.86 (s, 3H, CH3O). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 189.50, 163.29, 162.91, 155.49, 150.78, 137.91, 131.39, 131.12, 128.89, 127.88, 121.39, 113.72, 113.42, 111.12, 109.87, 100.58, 55.51. ESI-MS [m/z]: [M + H]<sup>+</sup> = 411.0; [M − H]<sup>−</sup> = 409.0. Anal. Calcd. for C19H14N4O3S<sup>2</sup> (%): C, 55.60; H, 3.44; N, 13.65; S, 15.62 Found (%): C, 55.52; H, 3.47; N, 13.73; S, 15.55.

#### *N***-{(5***Z***)-5-[(6-Methoxy-1***H***-indol-3-yl)methylene]-4-oxo-2-thioxothiazolidin-3-**

**yl}isonicotinamide 5k.** Yield 89%; m.p. 275–277 ◦C (CH3COOH). IR (cm–1): 3550.78 (NH), 3346.34 (NH), 1725.24 (C=O), 1689.56 (C=O), 1596.98 (C=C), 1576.73 (C=S). <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) δ 12.11 (s, 1H, NH), 11.85 (s, 1H, NH-CO), 8.78 (d, *J* = 5.3 Hz, 2H,H<sup>2</sup> +H6, pyridine), 8.11 (s, 1H, CH=), 7.89 (d, *J* = 5.3 Hz, 2H, H<sup>3</sup> +H5, pyridine), 7.74 (dd, *J* = 13.7, 5.6 Hz, 2H, H<sup>2</sup> +H4, indole), 6.96 (s, 1H, H<sup>7</sup> indole), 6.83 (d, *J* = 8.6 Hz, 1H, H<sup>5</sup> indole), 3.84 (s, 3H, CH3O). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 189.49, 163.30, 162.91, 156.97, 150.78, 137.88, 137.36, 130.66, 128.61, 121.38, 120.66, 119.52, 111.71, 111.17, 110.52, 95.46, 55.29. ESI-MS [m/z]: [M + H]<sup>+</sup> = 411.0; [M − H]<sup>−</sup> = 409.0. Anal. Calcd. for C19H14N4O3S<sup>2</sup> (%): C, 55.60; H, 3.44; N, 13.65; S, 15.62 Found (%): C, 55.73; H, 3.49; N, 13.57; S, 15.54.

**(5***Z***)-5-(1***H***-Indol-3-ylmethylene)-3-morpholin-4-yl-2-thioxothiazolidin-4-one7a.** Yield 86%; m.p. 273–275 ◦C (DMFA:CH3COOH). IR (cm–1): 3247.97 (NH), 1690.53 (C=O), 1594.09 (C=C), 1575.77 (C=S).1H-NMR (300 MHz, DMSO-d6, ppm) δ 12.10 (s, 1H, NH), 7.98 (s, 1H, CH=), 7.85 (d, *J* = 6.8 Hz, 1H, H<sup>4</sup> indole), 7.66 (d, *J* = 2.8 Hz, 1H, H<sup>7</sup> indole), 7.51–7.45 (m, 1H, H<sup>2</sup> indole), 7.27–7.13 (m, 2H, H<sup>5</sup> +H6, indole), 3.81 (s, 6H, morpholine), 3.06 (s, 2H, morpholine). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 189.92, 165.21, 136.32, 130.51, 126.72, 126.03, 123.34, 121.50, 118.48, 112.53, 111.68, 110.95, 66.56, 50.14. ESI-MS [m/z]: [M + H]<sup>+</sup> = 346.2; [M − H]<sup>−</sup> = 344.2. Anal. Calcd. for C16H15N3O2S<sup>2</sup> (%): C, 55.63; H, 4.38; N, 12.16; S, 18.56 Found (%):C, 55.74; H, 4.32; N, 12.24; S, 18.49.

(5*Z*)-5-[(1-Methyl-1*H*-indol-3-yl)methylene]-3-morpholin-4-yl-2-thioxo-thiazolidin-4-one 7b was prepared according to [42].

**(5***Z***)-5-[(5-Methoxy-1***H***-indol-3-yl)methylene]-3-morpholin-4-yl-2-thioxo-thiazolidin-4-one7c.** Yield 82%; m.p. 250–252 ◦C (CH3COOH). IR (cm–1): 3163.11 (NH), 1690.53 (C=O), 1580.59 (C=C, C=S). <sup>1</sup>H-NMR (300 MHz, DMSO-d6, ppm) δ 12.08 (s, 1H, NH), 7.99 (s, 1H, CH=), 7.61 (s, 1H, H<sup>4</sup> indole), 7.34 (d, *J* = 3.4 Hz, 2H, H<sup>2</sup> +H7, indole), 6.81 (d, *J* = 8.7 Hz, 1H, H<sup>6</sup> indole), 4.05–3.57 (m, 9H, CH3O, morpholine), 3.03 (s, 2H, morpholine). <sup>13</sup>C NMR (101 MHz, DMSO, ppm) δ 189.87, 165.20, 155.29, 131.04, 130.51, 127.75, 126.61, 113.52, 113.30, 111.04, 110.74, 100.34, 66.56, 55.46, 50.11. ESI-MS [m/z]: [M + H]<sup>+</sup> = 376.0; [M − H]<sup>−</sup> = 374.0. Anal. Calcd. for C17H17N3O3S<sup>2</sup> (%): C, 54.38; H, 4.56; N, 11.19; S, 17.08 Found (%):C, 54.31; H, 4.62; N, 11.04; S, 17.15.

#### *3.3. Antibacterial Activity Evaluation*

Bacterial strains utilized include Gram-negative: *Salmonella typhimurium* (ATCC 13311) *Pseudomonas aeruginosa* (ATCC 27853), *Escherichia coli* (ATCC 35210), *Enterobacter cloacae* (ATCC 35030) and Gram-positive bacteria: *Micrococcus flavus* (ATCC 10240), *Bacillus cereus* (isolated clinically), *Staphylococcus aureus* (ATCC 6538), and *Listeria monocytogenes* (NCTC 7973) bacteria. Pathogens were provided from the Mycological Laboratory, Institute for Biological Research "Siniša Stankovic" National institute of Republic of Serbia Belgrade. Resistant strains used were MRSA, *E. coli*, and *P. aeruginosa* [77,78].

For the determination of minimum inhibitory (MIC) and minimum bactericidal concentrations, the microdilution method, as previously described [77–79]. The minimum inhibitory (MIC) and minimum bactericidal (MBC) concentrations were determined by the modified microdilution method as previously reported [77–79]. Briefly, the fresh overnight culture of bacteria was adjusted to a concentration of 1 × 10<sup>5</sup> CFU/mL. The tested compounds were dissolved in 5% DMSO and serially diluted in tryptic soy broth (TSB) medium with bacterial inoculum (1.0 × 10<sup>4</sup> CFU per well). The microplates were incubated for 24 h at 37 ◦C. The MIC of the samples was detected following the addition of 40 µL of iodonitrotetrazolium chloride (INT) (0.2 mg/mL) and incubation at 37 ◦C for 30 min. The lowest concentration that produced a significant inhibition of the growth of the bacteria in comparison with the positive control was identified as the MIC. MBC was determined by serial sub-cultivation of 10 µL into microplates containing 100 µL of TSB. The lowest concentration that shows no growth after this sub-culturing was identified as the MBC, indicating 99.5% death of the original inoculum. Streptomycin and ampicillin were used as positive controls.

#### *3.4. Antifungal Evaluation*

The following fungi were used: Aspergillus niger (ATCC 6275), Aspergillus ochraceus (ATCC 12066), Aspergillus fumigatus (human isolate), Aspergillus versicolor (ATCC 11730), Penicillium funiculosum (ATCC 36839), Penicillium ochrochloron (ATCC 9112), Trichoderma viride (IAM 5061), Penicillium verrucosum var. cyclopium (food isolate). The organisms were obtained from the Mycological Laboratory, Department of Plant Physiology, Institute for Biological Research "Siniša Stankovic", National institute of Republic of Serbia, Belgrade, Serbia. All experiments were performed in duplicate and repeated three times, as previously described [80,81].

The fungal spores were washed from the surface of agar plates with sterile 0.85% saline containing 0.1% Tween 80 (*v*/*v*). The spore suspension was adjusted with sterile saline to a concentration of approximately 1.0 × 10<sup>5</sup> in a final volume of 100 µL per well. MIC determinations were performed by a serial dilution technique using 96-well microtiter plates. The examined compounds were serially diluted in broth Malt medium (MA), after which inoculum was added. The microplates were incubated for 72 h at 28 ◦C. The lowest concentrations without visible growth (at the binocular microscope) were defined as MICs. The fungicidal concentrations (MFCs) were determined by serial subcultivation of 2 µL of tested fractions dissolved in medium and inoculation into microtiter plates containing 100 µL of broth per well and further incubation 72 h at 28 ◦C. The lowest concentration with no visible growth was defined as MFC, indicating 99.5% killing of the original inoculum. The fungicides bifonazole and ketoconazole were used as positive controls.

#### *3.5. Docking Studies*

The program AutoDock 4.2® software was used for the docking simulation. The free energy of binding (∆G) of *E. coli* DNA GyrB, Thymidylate kinase, *E. coli* MurA, *E. coli* primase, *E. coli* MurB, DNA topo IV, and CYP51 of *C*. *albicans,* in complex with the inhibitors were generated using this molecular docking program. The X-ray crystal structures data of all the enzymes used were obtained from the Protein Data Bank (PDB ID: 1KZN, AQGG, 1DDE, JV4T, 2Q85, 1S16, and 5V5Z respectively). All procedures were performed according to our previous paper [78].

#### *3.6. Cytotoxicity*

**HEK 293** cells were cultured in DMEM medium, supplemented with 10% fetal calf serum (Sigma Chemical Co., St. Louis, MO, USA), 50 µg/mL streptomycin (Sigma Chemical Co.), and 50 units/mL penicillin (Sigma Chemical Co.) in 5% CO2-containing humidified atmosphere at 37◦C. Since compound solutions contained DMSO, control cultures containing only DMSO at the final concentration obtained when the appropriate volume of compound solution was added were performed.

#### MTT Assay for Determination of Cell Viability

MTT assay based on the colorimetric measurement of formazan formed after reducing MTT by cellular NAD(P)H-dependent oxidoreductases was used to examine the cytotoxic activity of the compounds. Briefly, the cells were seeded into 96-well plates in 100 µL of complete culture medium at a concentration of 5,000 substrate-dependent cells per well and left incubated overnight as described above. The formulations to be tested (100 µL aliquots) were added to the culture medium at different concentrations and left incubated for 72 h. The MTT assay was performed following the manufacturer's recommendations and assessed using an EL ×800 absorbance reader (BioTek Instruments; Winooski, VT, USA).

#### **4. Conclusions**

Eleven 5-[(R-1*H*-indol-3-yl)methylene]-4-oxo-2-thioxo-thiazolidin-3-ylcarbamides **5a-k** and three 5-[(R-1*H*-indol-3-yl) methylene] -3-morpholin-4-yl-2-thioxothiazolidin-4-ones **7a-c** were designed, synthesized and evaluated in silico and experimentally for their antimicrobial action against panel of Gram positive, Gram negative bacteria and fungi.

It should be mentioned that all compounds appeared to be more potent than ampicillin against all bacteria tested and then streptomycin against all bacteria except *B. cereus (isolated clinically M. flavus* (ATCC 10240)*,* and *En. cloacae* (ATCC 35030). The most sensitive bacteria was found to be *S. aureus* (ATCC 6538), while *L. monocytogenes* (NCTC 7973) was the most resistant one. Compounds also appeared to be active against three resistant strains MRSA, *E. coli,* and *P. aeruginosa* showing better activity against MRSA than both reference drugs while against the other two resistant strains better than ampicillin.

Concerning antifungal action, the tested compounds exhibited very good activity against all the fungal species tested, being more active than ketoconazole and bifonazole. The most sensitive fungal strain appeared to be *T. viride* (IAM 5061), while the most resistant filamentous *A. fumigatus* (human isolate).

It can be observed that the growth of both Gram-negative and Gram-positive bacteria and fungi responded differently to the tested compounds, which indicates that different substituents may lead to different modes of action or that the metabolism of some bacteria/fungi was better able to overcome the effect of the compounds or adapt to it.

Docking analysis to DNA Gyrase, Thymidylate kinase and *E.coli* MurB indicated a probable involvement of MurB inhibition in the antibacterial mechanism of compounds tested while docking analysis to 14α-lanosterol demethylase (CYP51) and tetrahydrofolate reductase of *Candida albicans* indicated a likely implication of CYP51 reductase at the antifungal activity of the compounds and secondary involvement of dihydrofolate reductase inhibition at the mechanism of action of the most active compounds.

Since the most active compounds **5d**, **5g**, **5k**, **7c** demonstrated the low cytotoxicity against HEK-293 human embryonic kidney cell line and reasonable selectivity index, this chemical series looks promising for investigations as the antimicrobial agents.

Finally, compounds **5d** (Z)-N-(5-((1H-indol-3-yl)methylene)-4-oxo-2-thioxothiazolidin-3 -yl)-4-hydroxybenzamide and **5g** (Z)-N-(5-((5-methoxy-1H-indol-3-yl)methylene)-4-oxo-2 thioxothiazolidin-3-yl)nicotinamide as well as **7c** (Z)-5-((5-methoxy-1H-indol-3-yl)methylene)-3 morpholino-2-thioxothiazolidin-4-one can be considered as lead compounds for further development of more potent and safe antibacterial and antifungal agents.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/1424-8247/13/9/229/s1, Supplementary file PASSweb\_results\_13mols.xlsx: Predictions of antimicrobial activity and acute rat toxicity.

**Author Contributions:** Conceptualization, A.G. and V.K.; methodology, V.H.; software, P.A. and P.P.; formal analysis, V.M.; investigation, M.I., M.K. and M.D.S; data curation, A.G., V.P., M.D.S. and P.E., original draft preparation, A.G. and P.P.; review & editing, A.G. and V.P.; supervision, A.G. and V.P. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was supported by the Serbian Ministry of Education, Science and Technological Development for financial support (project No. 451-03-68/2020-14/200007).

**Acknowledgments:** Computational predictions of biological activity by AntiBac-Pred, AntiFun-Pred and AcuTox web-services (P.P. and V.P.) were performed in the framework of the Russian State Academies of Sciences Fundamental Research Program for 2013–2020.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Communication*

### **A Fluorinated Analogue of Marine Bisindole Alkaloid 2,2-Bis(6-bromo-1***H***-indol-3-yl)ethanamine as Potential Anti-Biofilm Agent and Antibiotic Adjuvant Against** *Staphylococcus aureus*

**Ra**ff**aella Campana <sup>1</sup> , Gianmarco Mangiaterra <sup>2</sup> , Mattia Tiboni <sup>1</sup> , Emanuela Frangipani <sup>1</sup> , Francesca Biavasco <sup>2</sup> , Simone Lucarini 1,\* and Barbara Citterio 1,\***


Received: 12 August 2020; Accepted: 25 August 2020; Published: 26 August 2020

**Abstract:** Methicillin resistant *Staphylococcus aureus* (MRSA) infections represent a major global healthcare problem. Therapeutic options are often limited by the ability of MRSA strains to grow as biofilms on medical devices, where antibiotic persistence and resistance is positively selected, leading to recurrent and chronic implant-associated infections. One strategy to circumvent these problems is the co-administration of adjuvants, which may prolong the efficacy of antibiotic treatments, by broadening their spectrum and lowering the required dosage. The marine bisindole alkaloid 2,2-bis(6-bromo-1*H*-indol-3-yl)ethanamine (**1**) and its fluorinated analogue (**2**) were tested for their potential use as antibiotic adjuvants and antibiofilm agents against *S. aureus* CH 10850 (MRSA) and *S. aureus* ATCC 29213 (MSSA). Both compounds showed antimicrobial activity and bisindole **2** enabled 256-fold reduction (ΣFICs = 0.5) in the minimum inhibitory concentration (MIC) of oxacillin for the clinical MRSA strain. In addition, these molecules inhibited biofilm formation of *S. aureus* strains, and compound **2** showed greater eradicating activity on preformed biofilm compared to **1**. None of the tested molecules exerted a viable but non-culturable cells (VBNC) inducing effect at their MIC values. Moreover, both compounds exhibited no hemolytic activity and a good stability in plasma, indicating a non-toxic profile, hence, in particular compound **2**, a potential for in vivo applications to restore antibiotic treatment against MRSA infections.

**Keywords:** MRSA; marine bisindole alkaloids; antibiofilm activity; adjuvants agents; VBNC cells

#### **1. Introduction**

The diffusion of methicillin resistant *Staphylococcus aureus* (MRSA) strains is considered among the most common causes of health care-associated infections (HAIs) in hospitalized people all around the world [1]. Treatment of MRSA infections is limited, mainly because these microorganisms can develop resistance to multiple antibiotics [2,3]. Moreover, MRSA infections can be worsened by the ability of MRSA strains to grow as matrix-enclosed communities called biofilms, which promote adhesion and favor long-term survival on both biotic and abiotic surfaces [4,5].

Another strategy that MRSA have developed to resist antibiotic treatment is the ability to enter a state of dormancy, either as viable but non-culturable cells (VBNC) or as antibiotic persisters (APs) [6,7]. The complex architecture of biofilms, presenting different stressful microenvironments, may lead to

a spatial-physiologic heterogeneity of the embedded bacterial population [8,9] and, in this peculiar survival structure, both VBNC cells and AP can be present. Considering that VBNC cells have been shown to tolerate a wide variety of stressors, including starvation, growth inhibiting temperatures, suboptimal salinity, suboptimal pH and antibiotics [10], their role in biofilm biology is of great concern in view of developing new therapeutic approaches. One of these includes the use of antibiotic adjuvants (i.e., molecules that have the potential to improve the effectiveness of an antibiotic against which bacteria have developed resistance), by reducing the bacterial resistance to it, hence prolonging the lifespan of life-saving drugs [11]. lead to a spatial-physiologic heterogeneity of the embedded bacterial population [8,9] and, in this peculiar survival structure, both VBNC cells and AP can be present. Considering that VBNC cells have been shown to tolerate a wide variety of stressors, including starvation, growth inhibiting temperatures, suboptimal salinity, suboptimal pH and antibiotics [10], their role in biofilm biology is of great concern in view of developing new therapeutic approaches. One of these includes the use of antibiotic adjuvants (i.e., molecules that have the potential to improve the effectiveness of an antibiotic against which bacteria have developed resistance), by reducing the bacterial resistance to it, hence prolonging the lifespan of life-saving drugs [11]. In this context, marine sessile organisms can represent a suitable source of bioactive products

In this context, marine sessile organisms can represent a suitable source of bioactive products that are normally used to contrast bacteria diffused in the surrounding water [12]. These molecules have shown activity and selectivity against a wide spectrum of pharmacological targets, and their structures are often used as leads in drug discovery and development [13]. Among the various structural classes, bisindole alkaloids have attracted the attention of many researchers for their biological activities [14–16], especially their antimicrobial and antibiofilm activities [17–19]. Most of them present bromine and/or chlorine substitutions but there are no known examples of natural marine alkaloids containing fluorine. One of the possible reasons of the absence of fluorometabolites could be due to the very low abundance of fluoride ion (1.3 ppm) in the oceans with respect to chloride (Cl<sup>−</sup> = 20,000 ppm) and bromide (Br<sup>−</sup> = 70 ppm). On the other hand, about 20–25% of drugs on the market contain at least one fluorine atom. The very special effects of fluorine are very difficult to fully rationalize and its high presence in anthropogenic bioactive molecules has generally arisen from intense structure–activity relationship studies [20]. However, some of the effects of fluorine substitution are relatively straightforward to interpret such as the ability of fluorinated molecules to suppress metabolism relative to their hydrocarbon analogues. Moreover, organo-fluorine compounds are biologically and chemically more stable than the corresponding chlorine and bromine containing compounds [21]. that are normally used to contrast bacteria diffused in the surrounding water [12]. These molecules have shown activity and selectivity against a wide spectrum of pharmacological targets, and their structures are often used as leads in drug discovery and development [13]. Among the various structural classes, bisindole alkaloids have attracted the attention of many researchers for their biological activities [14–16], especially their antimicrobial and antibiofilm activities [17–19]. Most of them present bromine and/or chlorine substitutions but there are no known examples of natural marine alkaloids containing fluorine. One of the possible reasons of the absence of fluorometabolites could be due to the very low abundance of fluoride ion (1.3 ppm) in the oceans with respect to chloride (Cl<sup>−</sup> = 20,000 ppm) and bromide (Br<sup>−</sup> = 70 ppm). On the other hand, about 20–25% of drugs on the market contain at least one fluorine atom. The very special effects of fluorine are very difficult to fully rationalize and its high presence in anthropogenic bioactive molecules has generally arisen from intense structure–activity relationship studies [20]. However, some of the effects of fluorine substitution are relatively straightforward to interpret such as the ability of fluorinated molecules to suppress metabolism relative to their hydrocarbon analogues. Moreover, organo-fluorine compounds are biologically and chemically more stable than the corresponding chlorine and bromine containing compounds [21].

As a part of our ongoing investigations of the biological activities and possible applications of the marine bisindole alkaloid 2,2-bis(6-bromo-1*H*-indol-3-yl)ethanamine **1** [14–18] and for all the above described reasons, in the present work we focus our studies on natural compound **1** and its fluorinated analogue **2** [18] (Figure 1) to determine their potential as antibiotic adjuvants and antibiofilm agents against methicillin-susceptible (MSSA) and methicillin-resistant (MRSA) *S. aureus*. As a part of our ongoing investigations of the biological activities and possible applications of the marine bisindole alkaloid 2,2-bis(6-bromo-1*H*-indol-3-yl)ethanamine **1** [14–18] and for all the above described reasons, in the present work we focus our studies on natural compound **1** and its fluorinated analogue **2** [18] (Figure 1) to determine their potential as antibiotic adjuvants and antibiofilm agents against methicillin-susceptible (MSSA) and methicillin-resistant (MRSA) *S. aureus*.

**Figure 1.** Marine bisindole alkaloid 2-bis(6-bromo-3-indolyl) ethylamine **1** and its fluorinated **Figure 1.** Marine bisindole alkaloid 2-bis(6-bromo-3-indolyl) ethylamine **1** and its fluorinated analogue **2**.

analogue **2**. The experimental design included four different steps: (i) preliminary determination of the minimum inhibitory concentration (MIC) of each compound; (ii) assessment of their antimicrobial activity in combination with an antibiotic; (iii) determination of their antibiofilm properties in terms of both inhibition of biofilm formation and disruption of preformed biofilms and (iv) investigation The experimental design included four different steps: (i) preliminary determination of the minimum inhibitory concentration (MIC) of each compound; (ii) assessment of their antimicrobial activity in combination with an antibiotic; (iii) determination of their antibiofilm properties in terms of both inhibition of biofilm formation and disruption of preformed biofilms and (iv) investigation of their activity against VBNC *S. aureus* forms.

#### of their activity against VBNC *S. aureus* forms. **2. Results**

#### **2. Results**  *2.1. Chemistry*

[18] and described in Scheme 1*.* 

*2.1. Chemistry*  Marine bisindole alkaloid **1** and fluorinated analogue **2** were synthesized as previously reported Marine bisindole alkaloid **1** and fluorinated analogue **2** were synthesized as previously reported [18] and described in Scheme 1.

*Pharmaceuticals* **2020**, *13*, x FOR PEER REVIEW 3 of 12

**Scheme 1.** Reaction conditions: (**a**) diphenyl phosphate, acetonitrile, 80 °C, 24 h and (**b**) K2CO3, methanol, reflux, 2 h. **Scheme 1.** Reaction conditions: (**a**) diphenyl phosphate, acetonitrile, 80 ◦C, 24 h and (**b**) K2CO<sup>3</sup> , methanol, reflux, 2 h.

#### *2.2. Antibacterial and Adjuvants Activities of Bisindoles 1 and* **2** *2.2. Antibacterial and Adjuvants Activities of Bisindoles 1 and 2*

The MIC of the compounds **1** and **2** against the MRSA (*S. aureus* CH 10850) and the MSSA (*S. aureus* ATCC 29213) strains is reported in Table 1, along with their hemolytic activity. In detail, the MIC of compound **1** was 2 µg/mL for both *S. aureus* strains, while compound **2** exhibited MIC values of 32 and 16 µg/mL against *S. aureus* CH 10850 and *S. aureus* ATCC 29213, respectively. The toxicity of both compounds toward mammalian cells, assessed by determining their ability to lyse human erythrocytes, resulted to be very low, with values of 2.96% ± 0.02% and 2.32% ± 0.06%, respectively*.*  The MIC of the compounds **1** and **2** against the MRSA (*S. aureus* CH 10850) and the MSSA (*S. aureus* ATCC 29213) strains is reported in Table 1, along with their hemolytic activity. In detail, the MIC of compound **1** was 2 µg/mL for both *S. aureus* strains, while compound **2** exhibited MIC values of 32 and 16 µg/mL against *S. aureus* CH 10850 and *S. aureus* ATCC 29213, respectively. The toxicity of both compounds toward mammalian cells, assessed by determining their ability to lyse human erythrocytes, resulted to be very low, with values of 2.96% ± 0.02% and 2.32% ± 0.06%, respectively.

**Table 1.** Antimicrobial activity (minimum inhibitory concentration (MIC), µg/mL) and hemolytic activity (%) of the examined compounds **1** and **2**. **Table 1.** Antimicrobial activity (minimum inhibitory concentration (MIC), µg/mL) and hemolytic activity (%) of the examined compounds **1** and **2**.


When tested in association with oxacillin against *S. aureus* CH 10850, compound **1** caused a MIC reduction from 256 to 128 µg/mL, with a ΣFICs = 1.0, indicating additivity (Table 2). More promising results were obtained with the combination oxacillin-compound **2**, that showed a MIC decrease of oxacillin (from 256 to 1 µg/mL) with ΣFICs of 0.5, indicating a synergistic effect (Table 2). When tested in association with oxacillin against *S. aureus* CH 10850, compound **1** caused a MIC reduction from 256 to 128 µg/mL, with a ΣFICs = 1.0, indicating additivity (Table 2). More promising results were obtained with the combination oxacillin-compound **2**, that showed a MIC decrease of oxacillin (from 256 to 1 µg/mL) with ΣFICs of 0.5, indicating a synergistic effect (Table 2).

**Table 2.** Reduction of oxacillin resistance in *S. aureus* CH 10850 (methicillin resistant *Staphylococcus aureus* (MRSA)) exerted by compound **1** and **2** and determination of the synergistic activity as assessed by checkerboard assay (ΣFICs). **Table 2.** Reduction of oxacillin resistance in *S. aureus* CH 10850 (methicillin resistant *Staphylococcus aureus* (MRSA)) exerted by compound **1** and **2** and determination of the synergistic activity as assessed by checkerboard assay (ΣFICs).


#### *2.3. Antibacterial Activity in Plasma 2.3. Antibacterial Activity in Plasma*

One of the main issues with indole and their derivatives is their instability in vivo due to oxidation [22,23]. Thus, the MIC of both compounds was evaluated after preincubation (0, 3 and 6 h) in 50% human plasma at 37 °C. MIC values of compound **2** were in line with what found in both *S. aureus* strains, while a 16-fold increase was observed for compound **1** (Figure 2). However, it is very remarkable that all antimicrobial activities remained constant even after 6 h of plasma preincubation, hence showing a good stability in the physiologically relevant time intervals (Figure 2). One of the main issues with indole and their derivatives is their instability in vivo due to oxidation [22,23]. Thus, the MIC of both compounds was evaluated after preincubation (0, 3 and 6 h) in 50% human plasma at 37 ◦C. MIC values of compound **2** were in line with what found in both *S. aureus* strains, while a 16-fold increase was observed for compound **1** (Figure 2). However, it is very remarkable that all antimicrobial activities remained constant even after 6 h of plasma preincubation, hence showing a good stability in the physiologically relevant time intervals (Figure 2).

*Pharmaceuticals* **2020**, *13*, x FOR PEER REVIEW 4 of 12

**Figure 2.** MIC of compounds **1** and **2** against *S. aureus* CH 10850 (**A**) and *S. aureus* ATCC 29213 (**B**), after preincubation in 50% blood plasma for 0, 3 and 6 h at 37 °C. Conservative estimates of three trials for each compound are shown. **Figure 2.** MIC of compounds **1** and **2** against *S. aureus* CH 10850 (**A**) and *S. aureus* ATCC 29213 (**B**), after preincubation in 50% blood plasma for 0, 3 and 6 h at 37 ◦C. Conservative estimates of three trials for each compound are shown.

#### *2.4. Antibiofilm Activity 2.4. Antibiofilm Activity*

Compound **1** was able to inhibit the biofilm formation of *S. aureus* CH 10850 and *S. aureus* ATCC 29213 at their MIC values (2 µg/mL), causing a biofilm reduction of 52.6% and 49.6%, respectively (Table 3). When tested at 2× MIC (4 µg/mL), no biofilm formation was observed in the wells after 24 h of incubation (100% of biofilm formation inhibition). Compound **2** at its MIC (32 µg/mL) was able to inhibit (61.0%) the biofilm formation of *S. aureus* CH 10850, reaching a 76.5% inhibition when used at 64 µg/mL (2× MIC). *S. aureus* ATCC 29213 biofilm formation, although less susceptible (58.9% inhibition) than *S. aureus* CH 10850 to the MIC (16 µg/mL) of compound **2**, was completely (100%) inhibited by 32 µg/mL (2× MIC) of the same compound (Table 3). Concerning the biofilm-disrupting ability, compound **1** successfully removed 37.5% of *S. aureus* CH 10850 biofilm and the 28.0% of *S. aureus* ATCC 29213 biofilm, after a 30 min treatment. Under the same experimental conditions, compound **2** exerted a greater disaggregating activity, removing 56.3% and 53.9% of preformed biofilm of *S. aureus* CH 10850 and *S. aureus* ATCC 29213 biofilms, respectively (Table 3). Compound **1** was able to inhibit the biofilm formation of *S. aureus* CH 10850 and *S. aureus* ATCC 29213 at their MIC values (2 µg/mL), causing a biofilm reduction of 52.6% and 49.6%, respectively (Table 3). When tested at 2× MIC (4 µg/mL), no biofilm formation was observed in the wells after 24 h of incubation (100% of biofilm formation inhibition). Compound **2** at its MIC (32 µg/mL) was able to inhibit (61.0%) the biofilm formation of *S. aureus* CH 10850, reaching a 76.5% inhibition when used at 64 µg/mL (2× MIC). *S. aureus* ATCC 29213 biofilm formation, although less susceptible (58.9% inhibition) than *S. aureus* CH 10850 to the MIC (16 µg/mL) of compound **2**, was completely (100%) inhibited by 32 µg/mL (2× MIC) of the same compound (Table 3). Concerning the biofilm-disrupting ability, compound **1** successfully removed 37.5% of *S. aureus* CH 10850 biofilm and the 28.0% of *S. aureus* ATCC 29213 biofilm, after a 30 min treatment. Under the same experimental conditions, compound **2** exerted a greater disaggregating activity, removing 56.3% and 53.9% of preformed biofilm of *S. aureus* CH 10850 and *S. aureus* ATCC 29213 biofilms, respectively (Table 3).

**Table 3.** Antibiofilm activities of compounds **1** and **2** against *S. aureus* CH 10850 and *S. aureus* ATCC 29213. **Table 3.** Antibiofilm activities of compounds **1** and **2** against *S. aureus* CH 10850 and *S. aureus* ATCC 29213.


CH 10850 (MRSA) 52.6% 100% 61.0% 76.5% 37.5% 56.3%

#### ATCC 29213 49.6% 100% 58.9% 100% 28.0% 53.9% *2.5. VBNC Forms Induction*

*2.5. VBNC Forms Induction*  The influence of compounds **1** and **2** on VBNC *S. aureus* cells induction was tested. Both strains exhibited an about 2log reduction of viable cells when exposed to 2× MIC of compound **2** (Figure 3). However only *S. aureus* 10850 showed a significant gap (corresponding to the VBNC amount) The influence of compounds **1** and **2** on VBNC *S. aureus* cells induction was tested. Both strains exhibited an about 2log reduction of viable cells when exposed to 2× MIC of compound **2** (Figure 3). However only *S. aureus* 10850 showed a significant gap (corresponding to the VBNC amount) between total viable and culturable cells. Compound **1** did not seem to induce VBNC forms, as demonstrated by the lack of discrepancy between culture and qPCR/flow cytometry (Figure 3).

between total viable and culturable cells. Compound **1** did not seem to induce VBNC forms, as demonstrated by the lack of discrepancy between culture and qPCR/flow cytometry (Figure 3).

**Figure 3.** Effects of compounds **1** and **2** on *S. aureus* cell populations in in vitro biofilms. Biofilms of both *S. aureus* CH10850 (**A**) and *S. aureus* ATCC 29213 (**B**), in the absence (CTL) or presence of the two compounds at concentrations 1× and 2× MIC were developed in vitro and the amount of total viable and culturable cells counted after 24 h incubation at 37 ◦C. The amount of culturable cells was determined by plate count (CFU) and that of total (i.e., culturable and non-culturable) viable cells by qPCR and/or flow cytometry (FC).

#### **3. Discussion**

Considering the wide spread of antimicrobial resistance, the search for new and more efficient therapeutic approaches against MRSA strains represents a priority. Several studies have evidenced that molecules based on indole scaffolds can be used against different bacterial species [18,24,25]. Recently, it has been demonstrated that selected marine-alkaloid-derived molecules possess adjuvant activity against multi drug resistant bacteria [19], thus opening new challenges in this field of investigation. Therefore, we tested the fluorinated bisindole **2**, as well as the lead natural compound **1**, toward two different—one susceptible and one resistant to methicillin—strains of *S. aureus* (MSSA ATCC 39213 and MRSA CH 10850, respectively). Both compounds showed an antimicrobial activity, although compound **1** resulted in being more efficient than its fluorinated analogue compound **2** (MIC, 2 vs. 32 µg/mL in the MRSA and MIC 2 vs. 16 µg/mL in the MSSA). Indeed, bisindoles are known to act as antimicrobials by two possible mechanisms of action: the positive charge on the nitrogen at physiological pH makes them cationic surfactants able to destabilize the cytoplasmic membrane. Moreover, they could possibly inhibit the bacterial pyruvate kinase as reported for similar bisindoles [26,27].

Our compounds were tested as possible adjuvants. Both restored oxacillin activity against the MRSA strain. Indeed, while tested in association with oxacillin, an additive effect (ΣFICs = 1.0) and a synergistic one (ΣFICs = 0.5), for compounds **1** and **2**, respectively, was found. Interestingly, the fluorine substitution in compound **2** was able to increase the adjuvant property of the natural

compound. In detail, the fluorinated bisindole **2** was able to decrease the oxacillin MIC by 256-fold (from 256 to 1 µg/mL) for the clinical MRSA strain studied here. To the best of our knowledge, an adjuvant property of this class of bisindoles has never been reported, hence, the data herein observed provide novel information and stress the potential use of these class of natural compounds, together with their synthetic analogues, to contrast antibiotic resistance. Based on these encouraging results, the toxicological aspect of both compounds was also investigated by assessing their ability to lyse human erythrocytes. Neither of the two compounds exhibited hemolytic activity, thus adding important information on their safety. Although a different behavior after plasma incubation was observed (i.e., 16-fold MIC increase of compound **1**), it is of note that both compounds retained their antimicrobial activities even after 6 h of preincubation in blood plasma, demonstrating their chemical stability to this body fluid at physiologically relevant time intervals.

Microbial biofilms generate serious human health problems, including infectious diseases such as endocarditis, periodontitis and bacteremia [4]. The obtained results evidenced the ability of the tested compounds to inhibit *S. aureus* biofilm formation, reaching in most cases the complete biofilm formation inhibition at 2× MIC concentration, as previously reported for similar compounds [18]. Only for *S. aureus* CH 10850 a lower biofilm formation inhibition was evidenced, stressing the higher resistance of MRSA to antimicrobials [4]. Considering the important role of indole in bacteria, the presence of two units of indole in both compounds **1** and **2** may suggest that the observed antibiofilm activity derives from a modulation of indole-based signaling pathways [18,28]. Indeed, it was found that intracellular indole and its derivatives can cause a temporary repression of the *agr*-quorum sensing system in *S. aureus* [29]. The eradication activity of compounds **1** and **2** was afterward assessed on preformed biofilms of *S. aureus* CH 10850 MRSA and *S. aureus* ATCC 39213. As shown, the fluorinated bisindole **2** had a most pronounced disaggregating activity (>50% for both the examined strains) compared to the natural compound **1** (maximum 37.5%). This result could be related to the presence of two fluorine atoms in compound **2**, conferring a reduced lipophilicity compared to the two bromine atoms of the natural product **1**. To possibly confirm this hypothesis, we calculated an important physicochemical property related to the lipophilicity/hydrophilicity of a molecule, the octanol–water portion coefficient (logP). LogP values are high for lipophilic molecules and low for hydrophilic ones. Calculated LogP (cLogP) were 2.36 for compound **2** and 3.68 for compound **1** (by OSIRIS Property Explorer) [30]. From the cLogP values, fluorinated derivative **2** is more than 10-fold less lipophilic than the natural product **1**, and therefore more soluble in the hydrophilic biofilm polysaccharide matrix, resulting in a more effective disaggregating activity on preformed biofilms.

Biofilm development can also lead to the induction of dormant cells, persistent bacterial phenotypes that seem to be suitable adjuvant targets [11]. Between the two tested molecules, compound **2** exerted a VBNC inducing effect at a 2× MIC concentration. Moreover, the bacterial response resulted strain-specific, as *S. aureus* ATCC 29213 exhibited a little difference between total viable and culturable cells (0.5 log), whereas *S. aureus* CH 10850 (MRSA) showed a four-log difference between qPCR/flow cytometry and CFU counts. This seems to indicate a more powerful action of compound **2** against this specific strain, as already suggested by the checkerboard assay, resulting in synergy. Indeed, this compound caused the same reduction of total viable cells in both strains, but CH 10850 exhibited a deeper state of dormancy (i.e., higher amount of VBNC forms) than the ATCC 29213 strain. These data confirm the role of methicillin resistance in the development of VBNC forms as a survival strategy to stress conditions [31]. Furthermore, the expression of the *mecA* gene, conferring methicillin resistance through PBP2a synthesis, has been suggested to correlate with a greater stability of VBNC forms of the same strain (*S. aureus* CH10850) under unfavorable conditions [32]. It is thus conceivable that compound **2** may exert antibiofilm activity even if it can constitute a stress factor able to induce VBNC *S. aureus* forms, as previously described for different antimicrobial compounds [31].

To briefly summarize, natural bisindole alkaloid **1** and its fluorinated derivative **2** showed antimicrobial activity against the tested *S. aureus* strains (MIC ranging from 2 to 32 µg/mL). Surprisingly, compound **2** (at 16 µg/mL) reduced the MIC of oxacillin from 256 to 1 µg/mL (ΣFICs = 0.5) for the clinical

MRSA strain. Although both molecules inhibited biofilm formation of *S. aureus* strains, compound **2** showed greater eradicating activity on preformed biofilm compared to the natural alkaloid **1**. None of the tested molecules exerted a VBNC inducing effect at their MIC values. Moreover, both compounds exhibited no hemolytic activity and a good stability in plasma, indicating a non-toxic profile. Although the tested compounds as well as the number of bacterial strains were limited, these preliminary results encourage us to further examine this class of interesting alkaloids, and in particular the fluorinated bisindole **2**, for in vivo applications to restore antibiotic treatment against MRSA infections.

#### **4. Materials and Methods**

#### *4.1. Chemistry*

All organic solvents used in this study were purchased from Sigma–Aldrich (St. Louis, MO, USA), Alfa Aesar (Haverhill, MA, USA), or TCI (Tokyo, Japan). Prior to use, acetonitrile was dried with molecular sieves with an effective pore diameter of 4 Å. Column chromatography purifications were performed under "flash" conditions using Merck (Darmstadt, Germany) 230–400 mesh silica gel. Analytical thin-layer chromatography (TLC) was carried out on Merck silica gel plates (silica gel 60 F254), which were visualized by exposure to ultraviolet light and an aqueous solution of cerium ammonium molybdate (CAM). ESI-MS spectra were recorded with a Waters (Milford, MA, USA) Micromass ZQ spectrometer. <sup>1</sup>H NMR and <sup>13</sup>C NMR spectra were recorded on a Bruker (Billerica, MA, USA) AC 400 or 100, respectively, spectrometer and analyzed using the TopSpin 1.3 (2013) software package. Chemical shifts were measured by using the central peak of the solvent.

#### 4.1.1. General Procedure for the Synthesis of Derivatives **1**–**2**

Diphenyl phosphate (0.02 mmol) was added to a solution of the appropriate indole derivative (0.4 mmol) and (trifluoroacetylamino)acetaldehyde dimethyl acetal (0.2 mmol) in anhydrous acetonitrile (0.2 mL), and the resulting mixture was stirred at 80 ◦C for 24 h in a sealed tube, monitoring the progress of the reaction by TLC and HPLC-MS. After cooling to room temperature, saturated aqueous NaHCO<sup>3</sup> (30 mL) and dichloromethane (30 mL) were added and the two phases were then separated. The aqueous solution was extracted with dichloromethane (3 × 20 mL). After drying over dry Na2SO4, the combined organic phases were concentrated in vacuum and the resulting crude product was utilized without further purification. A mixture of that crude trifluoroacetamide derivative and potassium carbonate (1 mmoL) in MeOH (1.87 mL) and H2O (0.13 mL) was stirred and heated at reflux for 2 h. The MeOH was removed under reduced pressure and water was added (30 mL). The aqueous solution was extracted with dichloromethane (3 × 30 mL) and the resulting solution was dried with Na2SO<sup>4</sup> and then concentrated in vacuum. The crude material was purified by flash chromatography on silica gel.

#### 4.1.2. 2,2-Bis(6-bromo-1*H*-indol-3-yl)ethanamine (**1**)

The physicochemical data of compound **1** are in agreement with those that were reported [18].

#### 4.1.3. 2,2-Bis(6-fluoro-1*H*-indol-3-yl)ethanamine (**2**)

Compound **2** was prepared employing 6-fluoro-1*H*-indole and was isolated by column chromatography (dichloromethane/methanol/ammonia, 95:4:1) as a white solid in 70% yield (two steps). TLC: Rf = 0.18 (silica gel; dichloromethane/methanol/triethylamine, 90:9:1; UV, CAM). MS (ESI): *m*/*z* 310 [M-H]−. <sup>1</sup>H NMR (400 MHz, CD3OD, 293 K): δ = 3.37–3.41 (m, 2H, CHC*H2*NH2), 4.52 (dd, 1H, *J*<sup>1</sup> = *J*<sup>2</sup> = 7.5 Hz, C*H*CH2NH2), 6.72 (ddd, 2H, *J*5−<sup>7</sup> = 2.0 Hz, *J*5−<sup>4</sup> = 9.0 Hz, *J*5−<sup>F</sup> = 9.5 Hz, H5), 7.04 (dd, 2H, *J*7−<sup>5</sup> = 2.0 Hz, *J*7−<sup>F</sup> = 9.5 Hz, H7), 7.14 (d, 2H, *J* = 3.0 Hz, H2), 7.44 (dd, 2H, *J*4−<sup>F</sup> = 5.0 Hz, *J*4−<sup>5</sup> = 9.0 Hz, H4) ppm. <sup>13</sup>C NMR (100 MHz, CD3OD, 293 K): δ = 37.2, 45.6, 96.7 (d, 2C, *J* = 26 Hz, C5), 106.5 (d, 2C, *J* = 24 Hz, C7), 116.4 (2C, C3), 119.5 (d, 2C, *J* = 10 Hz, C4), 122.3 (d, 2C, *J* = 3 Hz, C9), 123.6 (2C, C2), 137.0 (d, 2C, *J* = 12 Hz, C8), 159.7 (d, 2C, *J* = 233 Hz, C6) ppm. The main physicochemical data of compound **2** are in agreement with those published [18].

#### *4.2. Bacterial Strains*

*S. aureus* CH 10850 (MRSA) [33] and *S. aureus* ATCC 29213 (MSSA), belonging to the strain collection of the Department of Life and Environmental Sciences (DiSVA), Polytechnic University of Marche (Ancona, Italy), were used. All the strains were cultured in brain heart infusion (BHI) broth or agar (Oxoid, Basingstoke, UK), subcultured in mannitol salt agar (MSA; Oxoid) and stored at −80 ◦C in BHI broth supplemented with 20% glycerol.

#### *4.3. Determination of Minimum Inhibitory Concentration (MIC)*

The minimum inhibitory concentration (MIC) of each molecule was determined by the microdilution method [34], with minor modifications. Bacteria were grown for 6 h in BHI broth at 37 ◦C, then diluted in Mueller Hinton II (Oxoid) to obtain ca. 5 × 10<sup>5</sup> CFU/mL in 100 µL, in the presence of increasing concentrations (2–128 µg/mL) of each compound dissolved in molecular biology grade dimethyl sulfoxide (DMSO, Sigma). Positive and negative controls included MHB inoculated or not with bacterial suspensions, respectively. Preliminary assays were performed to exclude the possible bacteriostatic and/or bactericidal activity of the solvent (i.e., DMSO); in any case, the volume of DMSO never exceeded 5% (*v*/*v*) of the final total volume. Tetracycline was used as a reference antibiotic, for comparison. MIC was defined as the lowest concentration of compound able to inhibit bacterial growth after 24 h of incubation at 37 ◦C, as detected by the unaided eye. All the experiments were performed three times using independent cultures.

#### *4.4. Checkerboard Assays*

The synergy of the two compounds and oxacillin (Sigma–Aldrich, St. Louis, Missouri, USA) against MRSA *S. aureus* CH 10850 was evaluated by the checkerboard assays [35], performed using 2-fold increasing concentrations of both compound (from 8 to 0.125µg/mL for compound **1** and from 512 µg/mL to 8 µg/mL for compound **2**) and oxacillin (from 512 to 0.5 µg/mL). Since the two compounds were resuspended in DMSO, the upper limit of the concentrations range tested was determined considering a final concentration of 1% DMSO. The combinations of each compound and oxacillin were evaluated by fractional inhibitory concentration (FIC) index, interpreted as follows: ≤0.5, synergy; >0.5 and ≤1.0, additive; >1.0 and <4, indifferent and ≥4, antagonistic.

#### *4.5. Hemolytic Activity*

The hemolytic activity of both compounds was evaluated as described by Ghosh et al. [36]. Briefly, 4 mL of freshly drawn, heparinized human blood was diluted with 25 mL of phosphate buffered saline (PBS), pH 7.4. After washing three times in 25 mL of PBS, the pellet was resuspended in PBS to 20 vol %. A 100 µL amount of erythrocyte suspension was added to 100 µL of different concentrations of compounds **1** and **2,** respectively. PBS and 0.2% Triton X-100 were used as the negative and positive control, respectively. Each condition was tested in triplicate. After 1 h of incubation at 37 ◦C each well was centrifuged at 1200 × *g* for 15 min, the supernatant was diluted 1:3 in PBS and transferred to a new plate. The OD<sup>350</sup> was determined using the Synergy HT microplate reader spectrophotometer (BioTek, Winooski, VT, USA). The hemolysis (%) was determined as follows:

$$[(A - A\_0)(A\_{\text{total}} - A\_0)] \times 100\tag{1}$$

where *A* is the absorbance of the test well, *A*<sup>0</sup> the absorbance of the negative control, and *Atotal* the absorbance of the positive control; the mean value of three replicates was recorded.

#### *4.6. Plasma Stability Assay*

*S. aureus* CH 10850 and *S. aureus* ATCC 29213 were grown for 6 h in BHI broth and diluted in Mueller Hinton II to obtain a final concentration of 1.5 × 10<sup>6</sup> CFU/mL. Fresh human blood was centrifuged at 3000 rpm for 5 min to separate the plasma from the cells. Three aliquots of compound **1** and compound **2** were dissolved in DMSO at a concentration of 128 and 1024 µg/mL, respectively and diluted 2-fold in plasma to reach the final concentration of 64 and 512 µg/mL. After incubation at 37 ◦C for 0, 3 and 6 h [36], 50 µL of each compound serially diluted 1:2 in MHB were added to a 96-well plate containing 50 µL of bacterial suspensions in MHB and incubated at 37 ◦C for 24 h. MIC values were determined as mentioned above [34]. No change in MIC values among the trials performed after different plasma-preincubation times was considered a proof of plasma-stability.

#### *4.7. Biofilm Formation Inhibition*

Biofilms were developed in 24-well polystyrene plates (VWR). *S. aureus* strains were grown in tryptic soy broth (TSB, VWR, Radnor, PA, USA) at 37 ◦C for 24 h. The bacterial concentration was adjusted to 5 × 10<sup>6</sup> CFU/mL, as previously described, and 100 µL of each bacterial suspension were inoculated in 24-well polystyrene plates supplemented with the corresponding amount of the selected compounds at their MIC and 2× MIC values. Two wells were inoculated with bacteria in TSB, as controls. After 24 h of incubation at 37 ◦C, the wells were washed with PBS to eliminate unattached cells and covered with 0.1% (*v*/*v*) crystal violet (CV) dissolved in H2O for 15 min and then washed in PBS and air-dried. The remaining CV was dissolved in 85% ethanol for 15 min at room temperature and 200 µL from each well was transferred to a 96-well plate for spectrophotometric quantification at 570 nm (Multiscan Ex Microplate Reader, Thermo Scientific, Waltham, MA, USA). Each data point was averaged from at least 8 replicate wells. All assays were performed in triplicate using independent cultures.

#### *4.8. Biofilm-Disrupting Activity*

Biofilms of each *S. aureus* strain were prepared with the procedure described above. After 24 h of incubation at 37 ◦C, the biofilms were gently washed in PBS, covered with the right amount of each compound at its MIC value, and left in contact for 30 min. For each plate, two wells were treated with saline and used as negative controls. After treatment, the biomass was evaluated by CV staining as described above. All data were expressed as the mean of three independent experiments performed in duplicate.

#### *4.9. VBNC Detection*

To evaluate the induction of staphylococcal VBNC forms, *S. aureus* CH 10850 and *S. aureus* ATCC 29213 biofilms were developed in Petri dishes (Ø 35 mm) by inoculating OD<sup>650</sup> = 0.1 cultures in BHI broth, alone or supplemented with either compound **1** or compound **2** at their MIC and 2× MIC concentrations, and incubated at 37 ◦C for 24 h. At the end of the incubation, biofilms were gently washed with 1 mL of PBS to remove planktonic bacteria, detached and then resuspended in 1 mL of PBS.

#### 4.9.1. Culture-Based Detection of Staphylococci

To evaluate the amount of the culturable cells, ten-fold serial dilutions of each mechanically detached biofilm were performed. For all dilutions 100 µL were spread onto BHI agar plates, incubated at 37 ◦C for 24 h prior to the enumeration of CFU.

#### 4.9.2. Flow Cytometry Detection of Staphylococci

The abundance of total viable staphylococci, both culturable and non-culturable, was determined by flow cytometry. Assays were performed using 200 µL of a 1:1000 dilution of detached *S. aureus* biofilms after live/dead staining (1× SYBR Green and 40 µg/mL propidium iodide), in a Guava Millipore cytometer, and analyzed by the GUAVASOFT 2.2.3 software. To discriminate bacterial cells from the background, a gate for cell detection in side scatter and green fluorescence (GRN) was applied, using both channels at 488 nm and a threshold value in the GRN channel; SYBR green and propidium iodide fluorescence were excited using a 488 nm laser and collected at 525/30 and 617/30 nm, respectively. To better detect signals, they were logarithmically (4 decades) amplified and, to increase statistical significance, the total number of particles analyzed was set to 20,000 events/replicate. All assays were run in duplicate.

#### 4.9.3. qPCR Detection of Staphylococci

Total DNA was extracted from 1 mL of biofilm aliquots diluted 1:10 (0× and 1× MIC) or undiluted (2× MIC) in PBS. Aliquots were centrifuged at 16,000 × *g* for 7 min, resuspended in 1 mL of STE (Tris-HCl 10 mM, NaCl 100 mM EDTA 1 mM) buffer supplemented with sucrose 20%, lysozyme 2.5 mg/mL and lysostaphin 100 µg/mL, and incubated for 1 h at 37 ◦C. Then, each aliquot was centrifuged, resuspended in 100 µL of PBS and the DNA was extracted using the QiaAmp DNA kit (Qiagen, Venlo, The Netherlands) according to the manufacturer instructions; a final elution volume of 80 µL was used.

*S. aureus* abundance was determined by *nuc*-qPCR using a Qiagen's Rotor-GeneQ MDx thermocycler, 0.2 µM of each primer [31] 10 µL of 2 × Rotor-Gene SYBR Green PCR master mix (Qiagen), and 2 µL DNA. Cycling conditions were 95 ◦C for 5 min, followed by 35 cycles of 95 ◦C for 10 s, 60 ◦C for 10 s and 72 ◦C for 10 s. A melting curve was obtained by ramping the temperature from 59 to 95 ◦C (0.5 ◦C/10 s) and analyzed with Qiagen's Rotor-GeneQ MDx software. DNA of *S. aureus* CH10850 and RNase-free water were used as positive and negative controls, respectively.

The number of viable *S. aureus* cells was determined as previously described [37,38]. Considering that *nuc* is a single copy gene [39] the amount of amplified DNA (ng) was divided for the weight (2.38928 × 10−<sup>10</sup> ng) of *nuc*; and divided by 2 (qPCR template volume) to obtain the number of *S. aureus* cells corresponding to 1 µL of DNA extract. Staphylococcal abundance/mL of the original sample was then calculated multiplying the number of bacterial cell/µL of DNA extract by 80 (undiluted samples) or 800 (1:10 diluted samples), respectively. Plate counts were compared with both qPCR and flow cytometry quantifications; any discrepancy > 0.5 log was considered to attest the presence of a VBNC *S. aureus* subpopulation.

#### **5. Conclusions**

Marine alkaloid **1** and its fluorinated derivative **2** showed antimicrobial activity and inhibited biofilm formation of *S. aureus* strains. Moreover, bisindole **2** showed greater disaggregating activity (up to 56% for the examined strains) on preformed biofilm compared to the natural alkaloid **1**. Interestingly, compound **2** enabled 256-fold reduction in the MIC of oxacillin for the clinical MRSA strain herein studied. These encouraging data for analogue **2**, together with the evidences of its safety and stability, could represent the first step toward validating the potential of employing this adjuvant to restore oxacillin efficacy against MRSA infections.

**Author Contributions:** Conceptualization, R.C., F.B., S.L. and B.C.; methodology, R.C., B.C. and S.L.; validation, F.B., S.L. and B.C.; formal analysis, F.B., S.L. and B.C.; investigation, R.C., G.M. and M.T.; resources, S.L. and B.C.; data curation, R.C., E.F., F.B. and B.C.; writing—original draft preparation, S.L. and B.C.; writing—review and editing, R.C., G.M., M.T., E.F. and F.B.; visualization, S.L.; supervision, S.L. and B.C.; project administration, S.L. and B.C.; funding acquisition, E.F., F.B. and S.L. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was partially supported by a grant from the Department of Biomolecular Sciences (DISB), University of Urbino Carlo Bo (DISB\_FRANGIPANI\_PROG\_SIC\_ALIMENTARE).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

1. Calfee, D.P. Trends in community versus health care-acquired methicillin-resistant *Staphylococcus aureus* infections. *Curr. Infect. Dis. Rep.* **2017**, *19*, 48. [CrossRef]


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Bactericidal and In Vitro Cytotoxicity of** *Moringa oleifera* **Seed Extract and Its Elemental Analysis Using Laser-Induced Breakdown Spectroscopy**

**Reem K. Aldakheel 1,2, Suriya Rehman 3,\* , Munirah A. Almessiere <sup>1</sup> , Firdos A. Khan <sup>4</sup> , Mohammed A. Gondal 5,\* , Ahmed Mostafa <sup>6</sup> and Abdulhadi Baykal <sup>7</sup>**


Received: 9 July 2020; Accepted: 7 August 2020; Published: 13 August 2020

**Abstract:** In the current study, we present the correlation between the capability of laser-induced breakdown spectroscopy (LIBS) to monitor the elemental compositions of plants and their biological effects. The selected plant, *Moringa oleifera*, is known to harbor various minerals and vitamins useful for human health and is a potential source for pharmaceutical interventions. From this standpoint, we assessed the antibacterial and in vitro cytotoxicity of the bioactive components present in *Moringa oleifera* seed (MOS) extract. Detailed elemental analyses of pellets of MOSs were performed via LIBS. Furthermore, the LIBS outcome was validated using gas chromatography–mass spectrometry (GC-MS). The LIBS signal was recorded, and the presence of the essential elements (Na, Ca, Se, K, Mg, Zn, P, S, Fe and Mn) in the MOSs were examined. The bactericidal efficacy of the alcoholic MOS extract was examined against *Escherichia coli (E. coli)* and *Staphylococcus aureus (S. aureus)* by agar well diffusion (AWD) assays and scanning electron microscopy (SEM), which depicted greater inhibition against Gram-positive bacteria. The validity and DNA nuclear morphology of human colorectal carcinoma cells (HCT-116) cells were evaluated via an MTT assay and DAPI staining. The MTT assay results manifested a profoundly inhibitory action of MOS extract on HCT116 cell growth. Additionally, MOS extracts produced inhibitory action in colon cancer cells (HCT-116), whereas no inhibitory action was seen using the same concentrations of MOS extract on HEK-293 cells (non-cancerous cells), suggesting that MOS extracts could be non-cytotoxic to normal cells. The antibacterial and anticancer potency of these MOS extracts could be due to the presence of various bioactive chemical complexes, such as ethyl ester and D-allose and hexadecenoic, oleic and palmitic acids, making them an ideal candidate for pharmaceutical research and applications.

**Keywords:** antibacterial; anticancer; GC-MS; LIBS; *Moringa oleifera*; seed extract

#### **1. Introduction**

Traditional herbalists throughout the globe have emphasized the role of plants used as remedies to treat different diseases, like inflammation and bacterial infections [1]. Enormous research is being carried out, where plants are being tested for active compounds that have antibacterial, antifungal and anticancer activities [2,3]. One such plant is *Moringa oleifera* Lam. (MOL), which originates mainly from Africa, Asia and South America [4,5]. The moringa leaves and fruits are considered as vegetables in the Philippines, Thailand, India and Pakistan [6]. Recently, sundry countries (Mexico, Caribbean islands, Hawaii and Cambodia) have begun planting it for its plentiful health benefits and nutritional value in addition to its medicinal importance [7,8]. *Moringa* fruits and leaves have various biological applications in addition to being enriched with vitamins, minerals, and proteins. The WHO recommends using MOL as a food because of its superior nutritional values for human health [6]. It is a well-known fact that *Moringa* plants (leaves and fruits) were grown as for skin sanitation, an energy source and also to relieve tension [8,9]. Innumerable reports have discovered that MOL has numerous significant merits, such as antioxidant, antimicrobial, anticancer, anti-inflammatory, antiulcer, antihypertensive, anti-urolithic, anti-asthmatic, antidiabetic, analgesic, anti-aging, diuretic, cardiovascular, hepatoprotective, hypoglycemic and immunomodulatory characteristics [10–13]. The hypotensive, antibacterial and anticancer efficacies of the MOL leaves and fruits are due to the presence of sundry distinctive chemical compounds (benzyl glucosinolate, niazimicin, benzyl iso-thiocyanate complexes and pterygospermin) in their structure [11]. Several methods have been improved to extract the initial contents from MOL plants for the production of food supplements and medicines (with natural organic components) and the determination of their other health benefits. Extraction procedures based on pressurized liquid, ultrasound, microwaves and supercritical fluid have been introduced [14–17].

Nevertheless, all extraction mechanisms experience several restrictions in terms of using a large amount of harmful organic solvents, sample production proceedings and cost [13]. To get better results, LIBS has emerged as an efficient approach for the identification and quantification of elemental compositions from different medicinal plant extractions. Over the past decade, the LIBS method has broadly been exploited for analyzing chemical elements existing in diverse kinds of specimens. This analytical method has distinguishing properties, like cost effectiveness, real-time measurement, sensitivity, rapidity and in situ elemental analysis [18–20]. Additionally, this type of analysis is easy, eco-friendly and is less complicated when preparing the sample. It is known that intensive sample treatments frequently produce erroneous results due to the presence of contaminants and loss-related effects. Thus, the LIBS technique can accurately disclose the identity of various medicinal plants in a scientific manner. In addition, most edible plants consist of proteins, vitamins and minerals, which possess immense benefits to health [21,22].

The purpose of the study is to analyze the presence of different elements in *Moringa oleifera* seed (MOS) extracts by using the LIBS approach and to examine the biological activities of MOS extracts by evaluating the antibacterial and anticancer potencies.

#### **2. Results**

#### *2.1. Qualitative Analysis of MOS Using LIBS*

The laser-induced breakdown spectroscopy (LIBS) spectra (Figure 1) of the MOS extract were recorded in the range of 200 to 800 nm. For the detection of the major elements, the spectra were recorded from different spots of the pellets and by scanning at a 50 nm wavelength range each time. In order to reduce the background noise and to enhance the LIBS signal intensity, LIBS parameters, including the delay times (delay between the incident laser pulse and the recording of spectra), the number of accumulations and laser energies, were optimized prior to the application of the LIBS setup for MOS sample analysis. The recorded LIBS spectra of the MOSs comprised various significant spectral peaks of the atomic and ionic lines (with varying intensities) related to the abundance of elements present in the tested MOS samples. Based on the NIST database, the recorded spectral lines were identified and classified in terms of the characteristic elements present in the MOSs, suggesting their significant role towards antibacterial and anticancer activities. Clearly, the measured LIBS spectra (Figure 1) exhibited the presence of vital elements such as Ca, K, Mg, P, S, Fe, Mn, Zn, Na and Se in MOS samples.

**Figure 1.** LIBS emission line spectra of various elements recorded in the wavelength range of 245–537 nm of MOS. The signature lines for different vital minerals present in MOSs are indicated in the figure.

Table 1 presents the LIBS signal intensities of the spectral transition lines corresponding to various detected elements in MOSs. In accordance with the Boltzmann distribution, the intensities of the LIBS spectral lines have a direct relationship with the elemental contents (concentration) present in MOSs [23,24]. This correlation was attained by considering the intensity ratio of the detected elemental lines with that of the C line taken as the reference (247.8 nm). The achieved intensity ratios of the elements Ca, K, Mg, P, S, Fe, Mn, Zn, Na and Se were in the ranges of 2.7–1.7, 1.2–1.4, 0.9, 1.1, 1.1, 1.1–0.9, 1.0, 0.9, 1.2 and 1.1, respectively, which were consistent with those reported in the literature [16].


**Table 1.** The detection of spectral lines of the different elements present in *MOS* by using our LIBS system.

#### *2.2. Volatile Content Analyses of MOS using GC-MS*

The GC-MS analyses of the *Moringa oleifera* seed (MOS) showed the presence of 114 volatile complexes with diverse chemical groups: fatty acids, esters, ketones, alcohols, aldehydes, and hydrocarbons. Table 2 shows the retention times and Figure 2 displays the percentage composition of all the identified chemical compounds in MOS. The main types were oleic acid (22.53%), 2-3-di-hydroxy-propyl (13.48%), 9-octa-decenoic acid (*Z*), 2-3-di-hydroxy-propyl ester (11.35%), docosenamide (6.04%), ethyl oleate (6.03%), 1-3-propanediol, 2-ethyl-2-(hydroxyl-methyl) (5.52%), oleic anhydride (3.96%), 2-propanone and 1-1-dimethoxy (3.86%). These MOSs contain fatty acids and their ester derivatives (65.45%), alcohols (9.4%), nitrogen compounds (9.09%), ketones (5.34%) and aldehydes (2.88%). To form esters, fatty acids and alcohols in plants may undergo esterification.

**Figure 2.** GC-MS chromatograph of MOS.




**Table 2.** *Cont*.


#### **Table 2.** *Cont*.

RT: Retention time in minutes.

#### *2.3.* In vitro *Cytotoxic Activity of MOS*

The cell viability of the MOS extract-treated HCT-116 cells was evaluated using an MTT assay after 48 h. The percentage of cell viability of the MOS extract-treated HEK-293 and HCT-116 cells was determined. The MTT assay examined the percentage of cell viability. The number of viable cells present in the control group was compared with the MOS-treated cells. In Figure 3, we show that cancer cells (control group, without MOS treatment) showed 100% cell viability, whereas cancer cells treated with MOS extract showed a significant decrease, which suggests that MOS extract could induce a significant drop in the viability of cancerous cells compared to control ones (without treatment using the MOS extract).

**Figure 3.** Effect of MOS extract on colon cancer cells (HCT-116) after treatment for 48 h with different concentrations. The average cell viability was calculated by MTT assay (\*\* *p* < 0.01).

The average viability of the MOS extract-treated cancer cells at various concentrations showed quite encouraging outcomes, with *p* < 0.01. The specificity of the MOS extract in the concentration range of 30 to 100 µg/mL on normal cells was inspected using MTT assays after treatment for 48 h (Figure 4). When MOS extract was tested on normal cells (HEK-293), we found no inhibitory action.

**Figure 4.** Effect of MOS extract on normal cells (HEK-293) after treatment for 48 h treated different concentrations. The average cell viability was calculated by MTT assay.

The MOS extract showed content-dependent specificity when the data were taken from three replications. A Student's *t*-test was used to understand the difference between the two treated groups and the results are presented as the mean ± standard deviation (SD).

#### *2.4. Nuclear Breakdown of MOS Extract-Treated Cancerous Cells*

Figure 5 illustrates the MOS extract-treated and untreated cell morphology that was imaged using confocal scanning microscopy (CSM). The MOS extract-treated cancer cells exhibited stronger inhibitory action (Figure 5A) than the control sample (Figure 5B). The CMS images of the nuclear cell morphology of both control (untreated) and MOS extract-treated (66 µg/mL) samples after stained by DAPI showed a substantial loss (nuclear disintegration) because of the treatments.

**Figure 5.** Confocal staining by DAPI. (**A**) The HCT-116 cells (non-treated) and (**B**) HCT-116 cells treated with MOS (66 µg/mL), 200× magnifications.

It was deduced that the MOS extract has strong anti-cancerous activities in colon cancer cells (as supported by the GC-MS analysis).

#### *2.5. Antibacterial E*ffi*cacy of MOS Extract*

The antibacterial potency of the MOS extract was assessed on Gram-negative and Gram-positive bacteria using the AWD method, wherein the inhibited areas due to antibacterial action around the inoculated wells were measured. This zone of inhibition was produced by the diffusion of the active chemical constituents present in the MOS extract. These results confirm the impact of the MOS extract on *S. aureus* and *E. coli*. Interestingly, the MOS extract was found to produce better antibacterial action in *S. aureus* bacteria than *E. coli*.

In Figure 6, the MOS extract shows concentration-dependent inhibition of *S. aureus* bacteria, with inhibition zones ranging from 18 to 24 mm. On the other hand, the inhibition zones of *E. coli* were found to range from 6 to 20 mm. The maximum and minimum inhibition zone diameters were acquired with the corresponding extract contents of 250 and 50 µg/mL (Figures 6 and 7). No inhibition was observed in *E.coli* with 50 µg/mL, whereas a higher concentration showed inhibition of the bacteria.

**Figure 6.** Agar well diffusion (AWD) plates showing the inhibition zones. (**A**) *S. aureus* and (**B**) *E. coli.* **1**: 50 µg/mL, **2**: 100 µg/mL, **3**: 150 µg/mL, **4**: 200 µg/mL, **5**: 250 µg/mL of MOS, **6**: Control (DH2O).

**Figure 7.** The graphical illustration of the zones of inhibition in millimeters (mm) examined in both *S. aureus* and *E. coli* after treatment using different concentrations of MOS extract. Data are the means ± SD of three different experiments.

The morphological changes in bacteria treated with MOS were studied using SEM. The untreated cells of *E. coli* were seen as regular rod-shaped cells with smooth cell surfaces (Figure 8a). The *E. coli* subjected to treatment with MOS appeared as damaged cells (Figure 8b) and the cell number also showed a reduction with a significant alteration of the cell wall and membrane. Similarly, the treatment of *S. aureus* with MOS extract also showed significant morphological changes in the structure and number of cells (Figure 8d), whereas the control *S. aureus* cells appeared as regular cocci with intact cell surfaces (Figure 8c). The damaged cells lost their cellular integrity, which led to the death of bacterial cells.

**Figure 8.** SEM micrographs for the study of morphogenesis by MOS extract. (**a**) *E. coli* cells (non-treated) and (**b**) treated *E. coli* cells, (**c**) *S. aureus* cells (non-treated) and (**d**) treated *S. aureus* cells.

#### **3. Discussion**

The LIBS and GC-MS techniques were utilized for the identification and quantitation of the elements existing in the MOS extract. For the first time, the anticancer and antibacterial activities of the MOS extract were assessed. Our results show the presence of diverse elements in MOSs, as confirmed by different methods. The GC-MS results of the MOS extract verified the existence of diverse anticancer and antimicrobial compounds. The spectra from different points on the surface of MOS pellets were examined by the LIBS method. The outcomes proved that MOSs are rich in different minerals that are useful to humans as food and medicine. Besides, the seeds have an influential role in antibacterial activity because of the availability of the following elements: K, Fe, P, Ca, Mg, S, Mn, Na, Se and Zn. It is clear that the amounts of Ca, K and Mg present in the MOSs were greater than the other elements. These results give evidence that MOSs are rich in different minerals, which are highly useful to humans as food supplements and medicine, indicating their remarkable impact in regulating the level of blood pressure, blood lipids, regulating the stomach function, protecting the liver, strengthening the bones, generating protein and enhancing the immunity of the human body [23–25]. Furthermore, the existence of Se in MOSs plays a vital role by protecting from fatal diseases like cancer, cardiovascular disease, cognitive decline, and thyroid disease. Moreover, these seeds exhibit a powerful antibacterial activity due to the presence of the detected elements, which is evidenced clearly by the antimicrobial studies conducted in this work.

The analysis of GC-MS of the MOSs showed the presence of 114 volatile complexes with diverse chemical groups: fatty acids, esters, ketones, alcohols, aldehydes, and hydrocarbons. To form esters, fatty acids and alcohols in plants may undergo esterification [19]. It has been reported that most of these compounds possess anticancer activities. In addition, palmitic acid shows selective cytotoxicity against human leukemic cells. The fatty acids also possess both antifungal and antibacterial activities. The omega-9 fatty acid was a detected primary complex (oleic acid) in the MOSs, which has numerous human health benefits (preventing ulcerative colitis and reducing blood pressure with remarkable antioxidant efficiency [26,27]). Therefore, GC-MS measurement proved the presence of numerous

fatty acids and their related esters (cis-9-hexadecenoic acid (palmitoleic), oleic acid, octadecanoic (stearic) acids, n-hexadecenoic acid) and alcohols. It was reported that most of these compounds have anticancer activity. For example, D-allose inhibits cancer cell growth at G1 phase [17]. Palmitic acid has selective cytotoxicity against leukemic cells in humans. Fatty acids also have antifungal and antibacterial effects [28].

In the present study, the biological activities of MOS were evaluated. The cell viability of the MOS extract-treated (for 48 h) HCT-116 (cancer) cells was assessed via MTT assay. The viability status of the extract-treated HEK-293 (normal) and HCT-116 cells was determined. It was inferred that the MOS extract has strong anticancer activity in colon cancer cells. However, few reports done on MOS extract have shown that the enhanced anticancer activity of the extract correlated with the occurrence of high oleic acid and fatty acid contents [21]. Previously, it has been shown that *Moringa* plants (leaves and fruits) have been used for various applications, such as antioxidant, antimicrobial, anticancer, anti-inflammatory, antiulcer, antihypertensive, anti-urolithic, antidiabetic, anti-asthmatic, anti-aging, analgesic, diuretic, cardiovascular, hepatoprotective, hypoglycemic and immunomodulatory uses [22]. As per our knowledge, there is no information on whether *Moringa* leaf extract or seed extract cause any differential response in cancer cells. Nevertheless, it would be interesting to do a comparative study where the effects of *Moringa* seed and *Moringa* leaf extracts are examined in cancer cells. We have used HEK-293 (human embryonic kidney) cells as normal cells to compare the MOS anticancer activity to human colon cancer cell line HCT-116. The purpose was to check whether MOS extracts produce any cytotoxic effects in normal cells or not. In our studies, we have found that MOS extracts produce inhibitory action in colon cancer cells (HCT-116), whereas no inhibitory action was found using the same concentrations of MOS in HEK-293 cells (non-cancerous cells), which suggests that MOS extracts could be non-cytotoxic to normal cells.

MOS has 114 volatile complexes with diverse chemical groups: fatty acids, esters, ketones, alcohols, aldehydes and hydrocarbons. The hypotensive, antibacterial and anticancer efficacies of the MO leaves and fruits are due to presence of sundry distinctive chemical compounds (benzyl glucosinolate, niazimicin, benzyl iso-thiocyanate complexes and pterygospermin) in their structure [6]. Several methods have been improved to extract the initial contents from MOL plants for the production of food supplements medicines (with natural organic components) and the determination of their other health benefits. While we do not know the molecular mechanism of the anticancer activities of MOS extract in cancer cells, the role of the caspase signaling pathway cannot be ruled out in the process of programmed cell death. It would be interesting to study the different caspases, such as caspase-3 and caspase-9 in MOS extract-induced cell death.

The antimicrobial potency of theMOS extract was assessed in the Gram-negative and Gram-positive bacteria via AWD, wherein the inhibited areas around the inoculated wells were measured. Due to the occurrence of varying cell components, there is a discrepancy in the bactericidal action of the MOS extract for the two different types of test bacteria [29].

The study of morphogenesis using SEM showed that MOS extract affects the cell wall at the initial stage, and later penetrates and accumulates at the surface membrane. This leads to an interruption in the metabolic activities of bacterial cells and initiates cell death [30]. The present study illustrates that the MOS extract causes damage to the microbial cell surface, thereby causing significant antibacterial activity. The penetration of the extract into the bacterial cell alters the membrane integrity by structural alterations, the loss of membrane proteins, etc.

Natural compounds like fatty acids, esters, ketones, alcohols, aldehydes and hydrocarbons have been reported in a variety of plants [31]. The mechanisms of the antibacterial activities of such compounds is linked to their high affinity towards lipids due to their hydrophobic characteristics. Their antibacterial actions are evidently related to this lipophilic nature and to the bacterial membrane structure [32]. This leads the compounds to penetrate the cellular membrane of the microbial cell, which enhances the fluidity and permeability of membrane, alters the topology of membrane proteins and inflicts disruption in the respiration chain [33,34].

The present study indicated the presence of phenolic compounds and these are reported to disrupt the cell membrane, leading to the inhibition of cell metabolism causing the leakage of cellular content [35]. It has been found that phenolics inhibit processes associated with the cell membrane, for example, electron transport, ions, protein translocation, phosphorylation and other enzyme-dependent reactions. Therefore, the disturbed permeability of the cytoplasmic membrane may lead to cell death [36]. The interaction of plant bioactive compounds with bacterial cell membranes results in the inhibition of several Gram-positive and Gram-negative bacteria [37]. It has also been indicated that Gram-positive bacteria are more susceptible to the antibacterial action of natural compounds like fatty acids, esters, ketones, alcohols, aldehydes and hydrocarbons, compared to Gram-negative bacteria [38]. This is also concordant with the current study, owing to the fact that Gram-negative bacteria have an outer layer surrounding their cell wall, limiting the access of hydrophobic compounds.

Therefore, MOSs can serve as an antibacterial component that can be further studied and recommended as a potential antimicrobial and anticancer therapy. This natural source can be further evaluated and upgraded for pharmaceutical application.

#### **4. Materials and Procedures**

#### *4.1. Seed Assembly and Extract Preparation*

All the extracts used in this study were prepared from good quality MOSs procured from a place where they are naturalized, Dammam in Saudi Arabia (originally imported from India). The seeds, without a shell, were crushed to obtain fine powder, followed by compression to get pellets. Figure 9a–c shows different steps of sample production for detailed analysis with the LIBS technique. Diversified ratios of the MOS powder between 5–25 g was mingled with 220 mL of ethanol and were stirred for 5 h, then the MOS solutions were filtered and put into a rotary evaporator to get a dry powder. The samples (MOSs) were subjected to GC-MS and anticancer activity and antibacterial activity tests.

**Figure 9.** Pictorial view of the pelletization of *Moringa oleifera* seed samples, showing: (**a**) as-purchased *Moringa oleifera* seeds, (**b**) *Moringa oleifera* seeds without coat, (**c**) pelletized seed sample.

### *4.2. LIBS Setup*

Figure 10 depicts the modified LIBS setup that was employed to determine the chemical constituents (elemental compositions) present in the MOS specimens. A quadrupled Q-switched Nd:YAG pulse laser (QUV-266-5) with an energy of 30 mJ, a pulse width of 8 ns, a 266 nm wavelength and a 20 Hz repetition rate was used with a UV convex lens of focal length 30 mm to focus the pulses onto the MOS pellets (which acted as the target) to ablate them. A plasma plume was generated when the target was ablated by the laser source. The emitted plasma was detected/collected using a fiber optic system positioned at 45◦ and the other end (500 mm) was connected to a spectrometer (Andor SR 500i-A) via grating with approximately 1200 lines per mm. An automated sample holder able to move across the plane was used to mount the pellet to avoid the formation of crusts on the surface of the target as a result of several laser pulses on the sample. The spectrum was recorded by an Intensified charge-coupled device (ICCD) camera (Model iStar 320T, 690 × 255 pixels with delay time setting at 300 ns) and data were transferred to an interfaced online computer (PC).

**Figure 10.** Schematic illustration of the LIBSsetup employed for the detection of vital elements present in MOS.

#### *4.3. GC-MS Measurements*

GC-MS (Shimadzu GC-2010 Plus) was applied for the analysis of ethanol MOS extracts that were provided by a split/splitless auto-injector (AOC-20i series) and coupled to a QP2010 Ultra single quadrupole instrument. The secession with GC was accomplished by using an Rxi-5MS fused silica capillary column (Restek, USA) with 30 m long, 0.25 mm wide and 100 µm thick films. The temperature was raised at a constant rate (5 ◦C/min) from the initial 60 ◦C (retained for 0.5 min) up to a final 280 ◦C (kept for 5 min). The temperature of the inlet was 270 ◦C (in the splitless mode) and helium (used as carrier gas of purity 99.99%) was blown through the MS transfer line (280 ◦C) at a 1 mL/min rate. The electron impact modes were used to operate the ion source (70 eV of energy at 250◦C) and the mass spectrum (full scan) was recorded in the range of 33–550 *m*/*z*. The spectrum values were gained by

regulating the GC-MS and via a GC-MS solution. The compounds of emanated volatiles were disclosed by means of the NIST-11 and WILEY-9 libraries and by calculating the relative area of each compound.

#### *4.4. Anticancer Activity of MOS Extract*

#### 4.4.1. In Vitro Cell Viability and Cell Culture Assay

As already mentioned, the anticancer activities of the MOS extract were evaluated by treating HCT-116 and normal (HEK-293) human cell lines. The cells were obtained from Dr. Khaldoon M. Alsamman, College of Applied Medical Science, Imam Abdulrahman Bin Faisal University, Dammam, Saudi Arabia. First, cells were cultured in 96 well-plates according to the earlier specified method. Dulbecco's modified Eagle's medium (DMEM), supplemented with other reagents (selenium chloride, fetal bovine serum, L-glutamine and antibiotics), was added to the plates for cell growth [29,30].

The cells were treated with varying concentrations of MOS extracts (30–100 µg/mL) for 48 h, whereas in the control, no MOS extract was added. Equal concentrations of a solvent, dimethyl sulfoxide (DMSO) were used in both the control and MOS-treated samples. After 48 h, an MTT assay was performed for 4 h (Molecules, Wellington, New Zealand). Afterward, the growth medium was removed from the plates and (DMSO) was added to every well so the MTT formed formazan crystal. Then, the wells containing the cell cultures were checked at a wavelength of 570 nm via a micro-plate reader (Bio-Rad Labs., Boston, MA, USA). Finally, the recorded readings were analyzed through inbuilt software (Version 5.0, GraphPad Prism) and the statistical significance was studied via ANOVA tests.

#### 4.4.2. Nuclear Staining via DAPI

The cells were treated with MOS extract (at 0.066 µg/mL) for 48 h and, in the control group, no MOS extract was added. The effects of MOS extract on cell nuclei were estimated with DAPI staining. Cold paraformaldehyde (4%) was used to pretreat the cells, which were then washed with 0.1% of Triton X-100 made from phosphate-buffered saline (PBS). At the same time, both the control and MOS- treated cells were stained using PBS with DAPI of 1.0 µg/mL concentration followed by rinsing with X-100 (0.1%). Cell morphologies of both cells (control and the treated) were analyzed *via* (CSM) (CSM-Zeiss, Frankfurt, Germany).

#### *4.5. Antimicrobial Activity Assessment of MOS Extract*

The effect of MOS extract on antibacterial efficacy at 50, 100, 150, 200 and 250 µg/mL against the *E. coli* ATCC35218 and *S. aureus* ATCC29213 strains was assessed using the AWD method. The inoculum was made from fresh bacteria grown overnight at 37 ◦C using nutrient broth (NB). This was followed by the preparation of inoculum to 0.5 McFarland standard and 100 µL of inoculum were disseminated over the surface of Mueller–Hinton agar (MHA) plates and dried in an aseptic environment. Next, the inoculated plates (6 mm) were punched using a sterile cork-borer. From the prepared extract, around 50 µL were added to the wells and further kept at 37 ◦C for an overnight incubation. Thereafter, the bacterial inhibition zone diameter over the entire well was recorded to evaluate the inhibition by the MOS extracts [29].

#### *4.6. Antimicrobial Activity Assessment of MOS Extract Using SEM*

The effect of MOS extract was additionally studied to investigate the structural damage caused to the selected bacteria by using SEM. The adjusted bacterial cell density, as described above, was treated with 250 µg/mL MOS for overnight incubation. The assay was carried out as per the protocol described by Rehman et.al [29].

#### *4.7. Statistical Analysis*

In the present study, cell viability data are presented as mean (±) standard deviation (SD), which were obtained from three independent experimental repeats. One-way ANOVA followed by Dunnett's post hoc test with GraphPad Prism software version 5.0 (GraphPad Software, Inc., La Jolla, CA, USA) was done for the statistical analysis. *p* < 0.05 was considered to indicate a statistically significant difference.

#### **5. Conclusions**

The LIBS and GC-MS techniques were used to identify and quantify the elemental compositions of MOS extract. The antibacterial and antiproliferative effectiveness of the MOS extracts was evaluated. The LIBS spectra revealed the presence of various nutritional elements in the MOSs that are important for health. The GC-MS analysis reconfirmed the presence of several bioactive compounds in the MOS extract. The MTT assay and DAPI staining showed a significant impact of the MOS extract on the inhibition of the growth of the HCT-116 cells and the insignificant inhibitory action of the extract on the HEK-293 cells, indicating the excellent specificity of the extracts towards the cancer cells. The MOS extracts showed strong antibacterial activity in terms of the growth inhibition and morphogenic changes against *S. aureus* compared to *E. coli,* owing to their cell wall differences. Therefore, it is established that MOS extract can be a prospective antibacterial and anticancer agent for functional pharmaceutical formulations.

**Author Contributions:** M.A.G. conceptualized and designed the study; R.K.A., M.A.A., S.R. and F.A.K. carried out the experiments and prepared all the figures; M.A.A., S.R., A.M. and F.A.K. wrote the manuscript; S.R., M.A.A., M.A.G., F.A.K. and A.B. revised and finalized the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **Products Derived from** *Buchenavia tetraphylla* **Leaves Have In Vitro Antioxidant Activity and Protect** *Tenebrio molitor* **Larvae against** *Escherichia coli***-Induced Injury**

**Tiago Fonseca Silva <sup>1</sup> , José Robson Neves Cavalcanti Filho 1,2 ,**

**Mariana Mirelle Lima Barreto Fonsêca 1,3, Natalia Medeiros dos Santos <sup>1</sup> , Ana Carolina Barbosa da Silva <sup>4</sup> , Adrielle Zagmignan 5,6, Afonso Gomes Abreu <sup>6</sup> , Ana Paula Sant'Anna da Silva <sup>1</sup> , Vera Lúcia de Menezes Lima <sup>1</sup> , Nicácio Henrique da Silva <sup>1</sup> , Lívia Macedo Dutra <sup>7</sup> , Jackson Roberto Guedes da Silva Almeida <sup>7</sup> , Márcia Vanusa da Silva <sup>1</sup> , Maria Tereza dos Santos Correia 1,**† **and Luís Cláudio Nascimento da Silva 4,6,\* ,**†


Received: 28 December 2019; Accepted: 29 February 2020; Published: 16 March 2020

**Abstract:** The relevance of oxidative stress in the pathogenesis of several diseases (including inflammatory disorders) has traditionally led to the search for new sources of antioxidant compounds. In this work, we report the selection of fractions with high antioxidant action from *B. tetraphylla* (BT) leaf extracts. *In vitro* methods (DPPH and ABTS assays; determination of phenolic and flavonoid contents) were used to select products derived from *B. tetraphylla* with high antioxidant action. Then, the samples with the highest potentials were evaluated in a model of injury based on the inoculation of a lethal dose of heat-inactivated *Escherichia coli* in *Tenebrio molitor* larvae. Due to its higher antioxidant properties, the methanolic extract (BTME) was chosen to be fractionated using Sephadex LH-20 column-based chromatography. Two fractions from BTME (BTFC and BTFD) were the most active fractions. Pre-treatment with these fractions protected larvae of *T. molitor* from the stress induced by inoculation of heat-inactivated *E. coli*. Similarly, BTFC and BTFD increased the lifespan of larvae infected with a lethal dose of enteroaggregative *E. coli* 042. NMR data indicated the presence of aliphatic compounds (terpenes, fatty acids, carbohydrates) and aromatic compounds (phenolic compounds). These findings suggested that products derived from *B. tetraphylla* leaves are promising candidates for the development of antioxidant and anti-infective agents able to treat oxidative-related dysfunctions.

**Keywords:** oxidative stress; natural products; medicinal plants; anti-infective agents; alternative infection models

#### **1. Introduction**

A substantial amount of evidence has indicated the key role of free radicals and reactive oxygen species (ROS) in the etiology of degenerative pathologies associated with aging (Parkinson's and Alzheimer's diseases), cancer, cardiovascular diseases, and diabetes [1–7]. Free radicals are highly reactive molecules characterized by having unpaired electrons in the last valence layer, thus becoming potent oxidizing agents [8,9]. These entities are produced as a result of normal cellular metabolism and play an important role in cell function and signaling; however, they can also damage important macromolecules (DNA, proteins, and lipids), thereby impairing cellular functions and leading to cell death [2,3,8].

Due to the reactivity of free radicals, organisms have developed an efficient antioxidant defense system formed by enzymes (such as superoxide dismutase and catalase) and proteins (glutathione reductase, thioredoxin) [8,9]. However, in many situations, this system cannot cope with the overproduction of reactive species, generating a so-called oxidative stress state, which is related to the clinical manifestations described above [1]. An alternative way of combatting the damage caused by free radicals is the use of exogenous substances collectively called antioxidants [4,8,10].

The antioxidants (natural or synthetic) can act through different mechanisms in the organism, such as: (i) direct neutralization of free radicals; (ii) expression of molecules from the host antioxidant defense systems; (iii) inhibition of oxidant enzymes (leading to reduction of free radicals' generation/propagation) [10–13]. These compounds can also be used in the food industry to maintain the physical-chemical quality of fruits, meat, and other foods [14–16]. Due to the side effects of synthetic antioxidants, those from natural sources are preferred; this context leads to a constant search for plant-derived compounds with this property [4,10–12,14–18].

In addition, antioxidant compounds have been related to the therapeutic properties of the species considered medicinal. In fact, substances with antioxidant actions have been detected in different products derived from plants (juices, teas, extracts, infusions) used in the treatment and prevention of diseases [10–12,14,17]. However, the antioxidant potential of some medicinal plants is still unexploited; and the neotropical species called *Buchenavia tetraphylla* (Aubl.) RA Howard (synonymy *Buchenavia capitata*; Combretoideae family) is a good example. This plant is distributed from Cuba (Central America) to southeastern Brazil (South America). *B. tetraphylla* is popularly known as "tanimbuca" in Brazil, where it has ethnomedicinal importance for communities in the northeast region [19]. It is also known to have a broad spectrum of antimicrobial activity, inhibiting bacteria, fungi, and virus [20–22]. Buchenavianine and two derivatives (O-demethylbuchenavianine, N,O-bis-(demethyl)buchenavianine) have been isolated from *B. tetraphylla*. These compounds are classified as flavoalkaloids with a piperidine moiety at carbon 8 [20,23].

The antioxidant potential of plant products has been traditionally characterized by in vitro methods; however, the biological relevance of these tests has been contested by several works [10,24,25]. In general, in vitro methods have been limited to sample prospection and compound isolation. Thus, the need to employ cell-based methods and in vivo models for a better understanding of the pharmacological action of a candidate as an antioxidant agent is evident [10]. In this work, in vitro methods were used to select products derived from *B. tetraphylla* with antioxidant action. Then, the samples with the highest potentials were evaluated in alternative models of stress based on *Tenebrio molitor* larvae inoculated with *Escherichia coli*.

#### **2. Results**

#### *2.1. Comparison of the Antioxidant Activity of Extracts Obtained from B. tetraphylla Leaves*

Initially, the phenolic and flavonoid contents were compared in different extracts of *B. tetraphylla* (BTHE: hexane extract; BTCE: chloroform extract; BTEE: ethyl acetate extract; BTME: methanolic extract) (Table 1). Among the extracts, BTME presented higher concentrations of both classes of compounds with values of 123.03 ± 1.51 mg of gallic acid equivalent (GAE) per mg of dry extract (mg GAE/mg) and 108.90 ± 0.07 mg of quercetin equivalent (QE) per mg of dry extract (mg QE/mg) (*p* < 0.05). A Pearson coefficient of 0.71 was found between the phenolic and flavonoid contents, indicating a strong correlation.

**Table 1.** Comparative analysis of total phenolic compounds, flavonoid content, and DPPH radical scavenging of the crude extracts from the leaves of *Buchenavia tetraphylla*.


Legend: BTHE: hexane extract; BTCE: chloroform extract; BTEE: ethyl acetate extract; BTME: methanolic extract. In each row, the values with significant differences (*p* < 0.05) are indicated by different superscript letters (a, b, c, d). The results are expressed as the mean ± standard deviation calculated from three independent assays performed in triplicate (*n* = *3*).

The antioxidant potential of the extracts and fractions was evaluated using the DPPH (2,2-diphenyl-1-picrylhydrazyl radical) and ABTS ((2,2-azino-bis (3-ethylbenzo-thiazoline-6-sulfonic acid) radical) methods. In the DPPH assay, the highest in vitro antioxidant activity was observed for BTME with an EC<sup>50</sup> (half maximal effective concentration) of 79.04 µg/mL (Table 1) and higher scavenging action in almost all tested concentrations in relation to other extracts (*p* < 0.001). The EC<sup>50</sup> found for Trolox was 44.10 µg/mL (positive control). Similarly, in the ABTS assay, a greater scavenging action was observed for BTME (approximately 100%; *p* < 0.05), followed by BTEE (approximately 50%) (Figure 1A). Strong correlations were also found between the levels of phenolic compounds and the scavenger actions towards DPPH (0.96) and ABTS radicals (0.89). The flavonoid contents were moderately correlated with DPPH scavenging (0.68) and strongly correlated with ABTS scavenging (0.95). The results for both antioxidant assays were strongly related (0.85).

μ

**Figure 1.** Comparative evaluation of antioxidant activity by the ABTS assay of the crude extracts (**A**) and fractions (**B**) of *Buchenavia tetraphylla* leaves. BTHE: hexane extract; BTCE: chloroform extract; BTEE: ethyl acetate extract; BTME: methanolic extract; BTFA: Fraction A; BTFB: Fraction B; BTFC: Fraction C; BTFD: Fraction D; BTFE: Fraction E; BTFF: Fraction F; BTFG: Fraction G; BTFH: Fraction H; BTFI: Fraction I. In each graph, values with significant differences (*p* < 0.05) are indicated by different superscript letters (a, b, c). The results are expressed as the mean ± standard deviation calculated from three independent assays performed in triplicate (*n* = *3*).

#### *2.2. Comparison of Phenolic and Flavonoid Content and Antioxidant Activity of Fractions Obtained from Methanolic Extract of B. tetraphylla Leaves*

‐ ‐ μ μ μ μ μ μ μ μ μ μ μ μ μ μ μ Since a higher level of antioxidant activity was observed in BTME, it was submitted to fractionation using Sephadex LH-20 column chromatography. A total of 9 non-repetitive fractions were obtained (BTFA to BTFI). Among them, the highest phenolic content was detected in BTFD (168.99 ± 2.22 µg GAE/µg), followed by BTFC (156.02 ± 4.51 µg GAE/µg), BTFG (127.62 ± 19.11 µg GAE/µg), and BTFI (110.15 ± 0.78 µg GAE/µg) (Table 2). Almost the same pattern was observed for the flavonoid content; in this case, BTFC had the highest values (68.26 ± 2.87 µg QE/µg) (*p* < 0.0001), followed by BTFD (56.01 ± 5.54 µg QE/µg), BTFG (45.27 ± 4.13 µg QE/µg), and BTFI (39.29 ± 2.89 µg QE/µg) (Table 2). A strong correlation (Pearson coefficient of 0.88) was observed among the concentration of phenolic and flavonoid compounds in the fractions obtained from BTME. Following, the antioxidant action of each fraction was investigated, and BTFC showed the highest activity against the DPPH radical (EC50: 50.41 µg/mL), followed by BTFD (EC50: 237.7641 µg/mL), BTFG (EC50: 294.38 µg/mL), and BTFI (EC50: 376.25 µg/mL). On the other hand, the fractions BTFB, BTFC, BTFD, BTFF, and BTFI scavenged approximately 80% of the ABTS radical (*p* < 0.05), and no statistical differences were observed between them (Figure 1B).

μ μ

μ μ


**Table 2.** Comparative analysis of total phenolic compounds and flavonoid content and DPPH radical scavenging of the crude extracts from leaves of *Buchenavia tetraphylla*.

**Legend:** BTFA-BTFI: Fractions obtained from *Buchenavia tetraphylla* methanolic extract. In each row, the values with significant differences (*p* < 0.05) are indicated by different superscript letters (a, b, c, d, e). The results are expressed as the mean±standard deviation calculated from three independent assays performed in triplicate (*n*=*3*).

#### *2.3. Evaluation of the Hemolytic E*ff*ects of Extracts and Fractions from Buchenavia tetraphylla Leaves*

Further, the hemolytic potential of each extract or fraction was evaluated using human erythrocytes (Figures 2 and 3). BTHE, BTCE, and BTEE induced toxic effects when tested at the highest concentrations (500 µg/mL and 1000 µg/mL) (Figure 2A–C). In contrast, it was observed that the BTME and its fractions did not induce significant hemolytic activity, even at the highest tested concentrations (Figures 2D and 3). μ μ

**Figure 2.** Hemolytic activity of the crude extracts of *Buchenavia tetraphylla* leaves. (**A**) BTHE (hexane extract); **(B**) BTCE (chloroform extract); (**C**) BTEE (ethyl acetate extract); (**D**) BTME (methanolic extract). \*\*\* Significant differences in relation to triton-X (*p* < 0.0001). The results are expressed as the mean ± standard deviation calculated from three independent assays performed in quadruplicate (*n* = *4*).

‐

‐

‐ **Figure 3.** Hemolytic activity of the fractions obtained from the methanolic extract of *Buchenavia tetraphylla* leaves. (**A**) BTFA (Fraction A); (**B**) BTFB (Fraction B); (**C**) BTFC (Fraction C); (**D**) BTFD (Fraction D); (**E**) BTFE (Fraction E); (**F**) BTFF (Fraction F); (**G**) BTFG (Fraction G); (**H**) BTFH (Fraction H); (**I**) BTFI (Fraction I). \*\*\* Significant differences in relation to triton-X (*p* < 0.0001). The results are expressed as the mean ± standard deviation calculated from three independent assays performed in quadruplicate (*n* = *4*).

#### *2.4. E*ff*ects of Extracts and Fractions from Buchenavia tetraphylla Leaves on the Survival of Tenebrio molitor larvae Submitted to Stress Induced by Heat-Killed Escherichia coli*

*‐* ‐ ‐ Based on the results presented in the above activities, we decided to evaluate the effects of the methanolic extract and its most active fractions (BTFC and BTFD) in a model of stress induced by heat-killed *E. coli* OP50 in *T. molitor* larvae. The heat treatment was used to ensure that larval death was not induced by bacterial growth; it was caused by the components present in the bacteria, such as lipopolysaccharide (LPS). Additionally, we used the nonpathogenic *E. coli* OP50 strain [26]. In this sense, at this point, we did not evaluate the antimicrobial action of the extract/fractions, but the ability of these to inhibit stress pathways induced by the presence of the bacteria. First, we evaluated the effects of several concentrations (measured at OD<sup>600</sup> (optical density at 600 nm)) of heat-killed *E. coli* OP50 (data not shown). The best results were obtained with the suspension at an OD600 of 0.7; this dose induced the death of 50% of the larvae after 15 h and of 90% after 30 h. These larvae presented typical myelination points related to stress induction in this organism. The pre-treatment with BTME at 10 mg/kg was able to inhibit animal death, with survival rates of 100%, 70%, and 50% after 15 h, 30 h, and 60 h of infection, respectively. The concentration of 20 mg/kg showed no significant protective action (Figure 4A).

The results obtained with BTFD were even more significant. The group treated with 20 mg/kg exhibited viability rates of 90% and 80% after 45 h and 60 h, respectively. Similar results were obtained with BTFD at 10 mg/kg, where viability remained at 80% after 45 h and 70% after 60 h (Figure 4C). Regarding BTFC, the best protective action was observed for the dose of 10 mg/kg. In this case, survival rates remained at 70% up to 45 h and ended at 60% (60 h). In the case of the 20 mg/kg dose, after 15 h, 80% of the larvae remained viable, and this rate progressively decreased to 50% (30 h), 40% (45 h), and reached 30% after 60 h (Figure 4B).

‐ ‐ ‐ ‐ ‐ ‐ ‐ **Figure 4.** Effects of methanolic extract of *Buchenavia tetraphylla* leaves (BTME) and its fractions (BTFC and BTFD) on the survival of *Tenebrio molitor* larvae challenged with heat-killed *Escherichia coli* OP50. (**A**) Effects of BTME on the survival of *T. molitor* larvae challenged with heat-killed *E. coli* OP50; (**B**) effects of BTFC on the survival of *T. molitor* larvae challenged with heat-killed *E. coli* OP50; (**C**) effects of BTFD on the survival of *T. molitor* larvae challenged with heat-killed *E. coli* OP50. The larvae (*n* = 10/group) were pre-treated with each sample (at 10 mg/kg or 20 mg/kg) 12 h prior to inoculation of heat-killed bacteria. Larvae treated with phosphate-saline buffer (PBS) or *E. coli* OP50 (BAC) were used as negative or positive controls, respectively. In this set of assays, larvae survival was recorded each 12 h. The experiment was repeated three times.

#### *2.5. E*ff*ects of Fractions from Buchenavia tetraphylla Leaves in the Lifespan of Tenebrio molitor larvae Infected by Enteroaggregative Escherichia coli*

We also evaluated the efficacy of BTFC and BTFD in a model of infection provoked by Enteroaggregative *E. coli* 042. The larvae were treated with each fraction 2 h after the infection. *E. coli* 042 killed most of the larvae in less than 24 h (median survival of one day). In contrast, the group treated with *B. tetraphylla* had median survivals of two days (Figure 5). It is important to highlight that these fractions did not have antimicrobial activity towards *E. coli*.

‐ **Figure 5.** Effects of fractions from methanolic extract of *Buchenavia tetraphylla* leaves (BTFC and BTFD) on the survival of *Tenebrio molitor* larvae challenged with *Escherichia coli* 042. (**A**) Effects of BTFC on the survival of *Tenebrio molitor* larvae challenged with *E. coli* 042. (**B**) Effects of BTFD on the survival of *Tenebrio molitor* larvae challenged with *E. coli* 042. The larvae *(n* = *10*/*group)* received a lethal dose of EAEC 042 and after 2 h were treated with fraction BTFC and BTFD (at 10 mg/kg or 20 mg/kg). Larvae treated with phosphate-saline buffer (PBS) or EAEC 042 were used as negative or positive controls, respectively. In this set of assays, larvae survival was recorded each 24 h. The experiment was repeated three times.

#### *2.6. Nuclear Magnetic Resonance Analysis of Buchenavia tetraphylla Leaves*

‐ δ The <sup>1</sup>H NMR analysis and two-dimensional experiments revealed similar profiles for BTFC and BTFD. The intense overlapping made difficult the identification of metabolites; however, it was possible to suggest the presence of some classes of compounds. The fractions showed signals in spectral regions related to aliphatic and aromatic compounds (Figure 6). The signals at δ 0.50–5.90 ppm attributed to an aliphatic compound (such as terpenoids, fatty acids, carbohydrates) were more expressive in comparison to the signals at δ 6.00–8.50 ppm related to aromatic compounds (such as phenolic compounds). This was observed mainly in the BTFD fraction.

δ

**Figure 6.** Representative <sup>1</sup>H NMR spectrum of the active fractions (BTFC and BTFD) obtained from the methanolic extract of *Buchenavia tetraphylla* leaves.

#### **3. Discussion**

The role of oxidative stress in the development of severe clinical conditions has promoted the search for new antioxidant agents from natural products [1,2,10,24,25]. Herein, we report the selection of high antioxidant fractions from *B. tetraphylla* leaf extracts using in vitro and in vivo approaches. Products derived from this plant have been shown as potential candidates for the development of new antimicrobial agents [20–22]; however, their antioxidant actions have not been properly addressed. The in vitro results obtained in this study showed that the methanolic extract (BTME) had a higher antioxidant activity, and this effect was correlated with its higher phenolic and flavonoid content. Among the fractions obtained from this extract, the best potentials were found for BTFC and BTFD (these fractions also exhibited greater levels of phenolic compounds). Furthermore, these agents were not toxic towards human erythrocytes.

‐ These samples were selected to be evaluated in a model of stress induced by heat-killed *E. coli* in *T. molitor* larvae. This insect has been used as a model organism in studies of microbial pathogenesis and drug development (antimicrobial, antivirulence, and immunomodulator agents) [27–29]. Several factors have supported the use of this animal. First, *T. molitor* is susceptible to pathogens such as *Candida albicans*, *E. coli*, and *Staphylococcus aureus*, which are able to persist within the infected insect, and cause changes in tissues, hemolymph, and phagocytes [30–32]. Second, the immune system of this insect has some known signaling pathways, such as the Toll pathway, the prophenoloxidase cascade, and the autophagy pathway [33–36].

‐ The antioxidant defense system of *T. molitor* is composed of several antioxidant and detoxifying enzymes such as superoxide dismutases, peroxidases, catalases, as well as tyrosinase, acetylcholinesterase, carboxylesterase, and glutathione S-transferase [37–39]. Previous studies have shown that during infection, the *T. molitor* larvae overproduce reactive species in response to the pathogen presence, leading to increased activity of several antioxidants and detoxifying enzymes that are correlated with larvae death [37,38,40].

‐ The ability of *T. molitor* to produce reactive species in response to deleterious stimulus makes this insect a potential model for the study of antioxidant substances. Despite these advantages, no study has exploited *T. molitor* larvae for the screening of plant-derived antioxidant compounds. The protective action of the agent is evidenced by the increased survival of the treated larvae compared to untreated larvae. This approach using larvae could bring more information than those traditionally used for antioxidant prospecting, based on the chemical interaction of compounds and without biological relevance [10,24].

Since *T. molitor* is a multicellular organism, this approach also presents advantages over those that use cells as some insights into the toxicity of the antioxidant agent can also be assessed. Furthermore, the use of this insect offers some advantages in relation to *Caenorhabditis elegans*, the invertebrate organism traditionally used for in vivo antioxidant evaluation [41,42], such as ease of handling and direct inoculation of the compound agent in the larvae. In our in vivo model of stress induced by heat-killed *E. coli* (OP50 strain), BTFD induced higher protective effects than BTME and BTFC. However, in our assays using the live EAEC strain, the fractions had similar results (both increasing the larvae median survival in one day). It is important to highlight that BTFC and BTFD did no display antimicrobial activity *in vitro* (data not shown), suggesting that their protective effects are related to the antioxidant properties. In this sense, we showed the efficacy of two fractions rich in antioxidants to reduce the deleterious effects of *E. coli*-induced injury in *T. molitor* larvae.

Previous works reported the predominant presence of flavonoids and alkaloids in *B. tetraphylla* and other species from the same genus [20,21,43,44]. In this present research, the most bioactive fractions (BTFC and BTFD) showed a similar chemical composition with the presence of aliphatic (terpenes, fatty acids, carbohydrates) and aromatic compounds (phenolic compounds). In general, the pharmacological potentials of some terpenes are associated with their antioxidant action [45,46]. These studies were reviewed by Gonzalez-Burgos and Gomez-Serranillos [45], who highlighted some structural features involved in the antioxidant action of each type of terpene.

Among the classes of compounds detected, phenolic compounds are the most usually related to antioxidant activity. The high antioxidant abilities of phenolic compounds are related to their phenolic hydroxyl groups that can donate hydrogen atom or transfer electrons, resulting in the scavenging of harmful free radicals (such as hydroxyl radicals). The aromatic groups present in the phenolic acids can also delocalize the unpaired electron [47,48]. According to Dai and Mumper [47], the flavonoids are able to perform electron transfer (mainly due to the ortho-dihydroxy structure on the B ring) and electron delocalization (the 2,3-double bond with a 4-oxo function in the C ring, which relocates from the B ring). The authors also highlighted the essential role of 3- and 5-hydroxyl groups with the 4-oxo function in A and C rings and 3-hydroxyl groups.

#### **4. Material and Methods**

#### *4.1. Collection and Extract Preparation*

Leaves of *B. tetraphylla* were collected in November 2013, in Catimbau National Park (Pernambuco, Brazil). The samples were processed according to the taxonomic techniques, identified, and deposited in the Herbarium of Agronomic Institute of Pernambuco (voucher: IPA 80349). The leaves of *B. tetraphylla* were subjected to drying at room temperature and then pulverized using a Macsalab mill (Model 200 LAB). This material was stored in a closed, dark container until used.

For extraction, 25 g of the powder were mixed with 100 mL of the first solvent (hexane; for BTHE) on a rotary shaker table (125 rpm) at 25 ◦C. After 72 h, the sample was filtered, and the extracted liquid was dried in a rotary evaporator (45 rpm) at 50 ◦C. The remaining leaf residue was further extracted with 100 mL of chloroform (BTCE), and the above procedure was repeated completely, subsequently performed with ethyl acetate (BTEE), and finally, with methanol (BTME).

#### *4.2. Fractionation of the Methanolic Extract*

The methanolic extract (2 g) was fractionated by chromatography using a column (80 cm × 2.5 cm) incorporated with Sephadex LH-20 (GE Healthcare®, Chicago, IL, USA; 60 cm high). The systems of eluents were based on the combination of methanol and ethyl acetate (as shown below). A total of 120 fractions (7 mL each) were obtained and analyzed by fluorescent black light (25W; 127V; Empalux®, Curitiba, Brazil) and thin layer chromatography (POLYGRAM® SIL G60/UV254; 20 × 20 cm; 0.20 mm; Macherey-Nagel®, Düren, Germany). After these procedures, fractions with similar phytochemical profiles were gathered, resulting in 9 different fractions: BTFA (Fractions 1–4), BTFB (Fractions 5–11), BTFC (Fractions 12–18), BTFD (Fractions 19–26), BTFE (Fractions 27–36), BTFF (Fractions 37–54), BTFG (Fractions 55–61), BTFH (Fractions 62–69), and BTFI (Fractions 70–120). The elution systems (methanol:ethyl acetate; *v*/*v*) for obtaining each fraction were: 7:3 for BTFA; 6:4 for BTFB, BTFC, BTFD, BTFE, BTFF; 5:5 for BTFG; 6:4 for BTFH and BTFI.

#### *4.3. Total Phenolic Content*

Obtaining the total phenolic compounds in crude extracts and fractions was performed using the Folin-Ciocalteu reagent [49]. Samples of each extract/fraction (200 µL at 1000 µg/mL) were mixed with 1.0 mL of Folin-Ciocalteu reagent, and 800 µL of 20% sodium carbonate were added after 3 min. The mixture was incubated at room temperature, protected from light, and allowed to stand for 2 h. The absorbance of the mixture was measured at 765 nm (GeneQuant 1300, GE Healthcare). The total phenolic content was calculated in mg of gallic acid equivalent (GAE) per mg of dry extract using a calibration curve obtained with gallic acid (y = 0.0043x + 0.0153; R<sup>2</sup> = 0.9932). The results were expressed as mean ± standard deviation calculated from three independent assays performed in triplicate (*n* = *3*).

#### *4.4. Flavonoid Content*

For flavonoid content, 100 µL (at 1000 µg/mL) of each sample were mixed with 100 µL of the reagent solution (2 g of aluminum chloride diluted in 2% ethanol solution). The mixture was incubated at room temperature and protected from light, and after 60 min, the absorbance was measured at 420 nm [50]. The content of flavonoids was calculated in mg of quercetin equivalent (QE) per mg of dry extract using a calibration curve constructed with standard quercetin solution (y = 0.004x + 0.0121; R<sup>2</sup> = 0.993). The results are expressed as the mean ± standard deviation calculated from three independent assays performed in triplicate (*n* = *3*).

#### *4.5. DPPH Assay*

An aliquot of 250 µL of 1 mM DPPH solution (2,2-diphenyl-1-picrylhydrazyl; Sigma-Aldrich) was added to 40 µL of different sample concentrations (31.25–1000 µg/mL) and homogenized. After 30 min, the absorbance was measured at 517 nm [51]. Trolox was used as the control compound. The DPPH sequestering activity was calculated using the formula below. The results were expressed as the mean ± standard deviation calculated from three independent assays performed in triplicate (*n* = *3*).

DPPH scavenging (%) = (Ac − As)/Ac × 100

where: Ac = absorbance control; As = sample absorbance

#### *4.6. ABTS Assay*

The ABTS (2,2-azino-bis (3-ethylbenzo-thiazoline-6-sulfonic acid)) radical cation was prepared 16 h prior to the assay by mixing 5 mL of the stock solution (7 mM) with 88 µL of the 140 mM potassium persulfate solution. Aliquots (20 µL) of each extract/fraction and 2 mL of the ABTS radical were mixed, and the absorbance of the solutions was monitored at 734 nm after 6, 15, 30, 45, 60, and 120 min, respectively [52]. Gallic acid was used as the positive control. The ABTS scavenging was calculated using the formula below. The results were expressed as the mean ± standard deviation calculated from three independent assays performed in triplicate (*n* = *3*).

ABTS scavenging (%) = (Ac − Aa)/Ac × 100

where Ac (control absorbance) and Aa (sample absorbance).

#### *4.7. Hemolytic Activity*

Blood (5–10 mL) was obtained from healthy, non-smoker volunteers by venipuncture, after signing a free informed consent form. Human erythrocytes from citrated blood were immediately isolated by centrifugation at 1500 rpm for 10 min. After removal of the plasma, the erythrocytes were washed three times with phosphate-buffered saline (PBS; pH 7.4), and then, a suspension of 1% erythrocytes was prepared as the same buffer. Following, an aliquot of 1.1 mL of erythrocyte suspension was mixed with 0.4 mL of each extract/fraction (concentration range: 125 to 1000 µg/mL). The negative control and positive control received 0.4 mL of PBS and Triton X, respectively. After 60 min of incubation at room temperature, the cells were centrifuged, and the supernatant was used to measure the absorbance of hemoglobin released at 540 nm [21]. The hemolytic activity was expressed in relation to the action of Triton X-100 and calculated using the formula below. The results were expressed as the mean ± standard deviation calculated from three independent assays performed in quadruplicate (*n* = *4*).

$$\text{Hemolytic activity (\%)} = [(\text{Aa} - \text{Ab}) \times 100] \text{(Ac} - \text{Ab)}$$

where: Aa is sample absorbance; Ab is the absorbance of the negative control; and Ac is the absorbance of the positive control.

#### *4.8. Toxicity Model Using Heat-Killed E. coli*

Larvae of *T. molitor* (~100 mg) were randomly allocated into groups (*n* = *10*). After anesthesia and disinfection, 10 µL of the most active samples (methanolic extract or fractions C and D; at 10 mg/kg or 20 mg/kg) were injected in the ventral membrane between the second and third abdominal segments (tail to the head) [29]. One hour after the sample inoculation, the larvae received 10 µL of heat-killed *E. coli* OP50 (optical density at 600 nm: 0.7). The viability of the larvae was evaluated after 15, 30, 45, and 60 h (by evaluation the lack of movement after mechanical stimulus). Larvae inoculated with the microorganism and treated with PBS were used as the negative control; while larvae that received two doses of PBS were the positive control. The experiment was performed in three independent assays.

#### *4.9. Infection Model Using Enteroaggregative E. coli*

Larvae (*n* = *10*) were prepared as described above and infected with 10 µL of enteroaggregative *E. coli* (EAEC) 042 (optical density at 600 nm: 0.1). After two hours, each animal received 10 µL of BTFC or BTFD (at 10 mg/kg or 20 mg/kg). Larvae inoculated with *E. coli* 042 and treated with PBS were used as the negative control; while larvae that received two doses of PBS were the positive control. The experiment was repeated three independent assays.

#### *4.10. Nuclear Magnetic Resonance Analysis*

The chemical characterization of the most active fractions (BTFC and BTFD) was performed by nuclear magnetic resonance (NMR) analysis. 1D and 2D NMR data were acquired at 298 K in DMSO-d<sup>6</sup> on a Bruker AVANCE III 400 NMR spectrometer operating at 9.4 T, observing <sup>1</sup>H and <sup>13</sup>C at 400 and 100 MHz, respectively. The NMR spectrometer was equipped with a 5 mm direct detection probe (BBO) with a *z*-gradient. One-bond (1H-13C HSQC) and long-range (1H-13C HMBC) NMR correlation experiments were optimized for average coupling constant <sup>1</sup> *J*(C,H) and LR*J*(C,H) of 140 and 8 Hz, respectively. All <sup>1</sup>H and <sup>13</sup>C NMR chemical shifts (δ) were given in ppm related to the TMS signal at 0.00 as an internal reference and the coupling constants (*J*) in Hz.

#### *4.11. Statistical Analysis*

The results were expressed as the mean ± standard deviation (SD). Statistical significance was determined by one-way ANOVA or two-way ANOVA followed by Tukey and Bonferroni tests. A *p*-value < 0.05 was considered statistically significant. Determination of EC<sup>50</sup> (half maximal effective

concentration) was performed by linear regression. Correlations were assessed using the Pearson coefficient. The larvae survival assays were analyzed using the Kaplan–Meier method to calculate survival fractions, and the log-rank test was used to compare survival curves.

#### **5. Conclusions**

In this study, the use of in vitro antioxidant assays allowed the selection of fractions from a methanolic extract with a high activity and low toxicity. The fractions (BTFC and BTFD) were able to extend the lifespan of *T. molitor* larvae submitted to stress induced by heat-killed *E. coli* significantly. The therapeutic treatment with these fractions had also positive effects on the infection induced by the pathogenic strain *E. coli* 042 (EAEC). <sup>1</sup>H NMR data indicated the presence of aliphatic (terpenes, fatty acids, carbohydrates) and aromatic compounds (phenolic compounds). These findings suggested that products derived from *B. tetraphylla* leaves are a promising candidate for the development of antioxidant agents able to treat the oxidative-related dysfunctions.

**Author Contributions:** T.F.S., M.V.d.S., M.T.d.S.C., and L.C.N.d.S. conceived of the study and performed the study design. T.F.S., V.L.d.M.L., N.H.d.S., J.R.G.d.S.A., M.V.d.S., M.T.d.S.C., and L.C.N.d.S. provided the reagents and equipment for all assays. J.R.N.C.F., M.M.L.B.F., N.M.d.S., and A.P.S.d.S. prepared the extracts and performed the in vivo and in vitro antioxidant assays. A.C.B.d.S., A.Z., and A.G.A. performed the anti-infective assays with *T. molitor*. J.R.G.d.S.A. and L.M.D. performed the chemical characterization of the extracts. All authors interpreted and discussed the results. T.F.S., J.R.G.d.S.A., L.M.D., A.C.B.d.S., M.T.d.S.C., and L.C.N.d.S. drafted and revised the manuscript. All authors approved the final version of this manuscript.

**Funding:** This work was funded by Fundação de Amparo à Ciência e Tecnologia de Pernambuco (FACEPE) Fundação de Amparo à Pesquisa e Desenvolvimento Científico do Maranhão (FAPEMA; BEPP-02241/18), Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES), and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq).

**Acknowledgments:** The authors thank the support given by the Laboratory of Natural Products and the Laboratory of Molecular Biology, Department of Biochemistry, of the Federal University of Pernambuco (UFPE;Universidade Federal de Oer).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **In Vitro Assessment of Antimicrobial, Antioxidant, and Cytotoxic Properties of Saccharin–Tetrazolyl and –Thiadiazolyl Derivatives: The Simple Dependence of the pH Value on Antimicrobial Activity**

**Luís M. T. Frija 1,\*, Epole Ntungwe <sup>2</sup> , Przemysław Sitarek <sup>3</sup> , Joana M. Andrade <sup>2</sup> , Monika Toma <sup>4</sup> , Tomasz Sliwi ´nski ´ <sup>4</sup> , Lília Cabral <sup>5</sup> , M. Lurdes S. Cristiano <sup>5</sup> , Patrícia Rijo 2,6,\* and Armando J. L. Pombeiro <sup>1</sup>**


Received: 4 October 2019; Accepted: 7 November 2019; Published: 12 November 2019

**Abstract:** The antimicrobial, antioxidant, and cytotoxic activities of a series of saccharin–tetrazolyl and –thiadiazolyl analogs were examined. The assessment of the antimicrobial properties of the referred-to molecules was completed through an evaluation of minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) values against Gram-positive and Gram-negative bacteria and yeasts. Scrutiny of the MIC and MBC values of the compounds at pH 4.0, 7.0, and 9.0 against four Gram-positive strains revealed high values for both the MIC and MBC at pH 4.0 (ranging from 0.98 to 125 µg/mL) and moderate values at pH 7.0 and 9.0, exposing strong antimicrobial activities in an acidic medium. An antioxidant activity analysis of the molecules was performed by using the DPPH (2,2-diphenyl-1-picrylhydrazyl) method, which showed high activity for the TSMT (*N*-(1-methyl-2*H*-tetrazol-5-yl)-*N*-(1,1-dioxo-1,2-benzisothiazol-3-yl) amine, **7**) derivative (90.29% compared to a butylated hydroxytoluene positive control of 61.96%). Besides, the general toxicity of the saccharin analogs was evaluated in an *Artemia salina* model, which displayed insignificant toxicity values. In turn, upon an assessment of cell viability, all of the compounds were found to be nontoxic in range concentrations of 0–100 µg/mL in H7PX glioma cells. The tested molecules have inspiring antimicrobial and antioxidant properties that represent potential core structures in the design of new drugs for the treatment of infectious diseases.

**Keywords:** saccharin; tetrazole; 1,3,4-thiadiazole; H7PX glioma cells; antimicrobial screening; antioxidant capacity

#### **1. Introduction**

In 1878, 1,2-benzisothiazole-3-one 1,1-dioxide (**1**, Figure 1), commercially known as saccharin, was discovered accidentally by Fahlberg during an investigation of the oxidation of *o*-toluenesulfonamide [1,2]: it was published by Remsen and Fahlberg one year later [3]. For more than a century, saccharin has been commonly used as a noncaloric artificial sweetener in the form of its water-soluble salts (mainly sodium, ammonium, and calcium), and it is the principal sweetening component of diabetic diets. For about three decades (since reports on carcinogenicity in laboratory animals were published), the debate on its toxicity to humans has not reached a consensus [4–6]. Numerous *N*-substituted derivatives of saccharin have been assessed for in vitro biological activity [7–10]. For example, first-row transition metal saccharinates as well as dioxovanadium(VI), dioxouranium(VI), and cerium(IV) saccharinates have been classified as protease inhibitors, and several metal(II) saccharinates have displayed superoxide dismutase-like activity [11]. Besides, structure–activity relationship studies have shown that the saccharin scaffold is an effective element for the development of inhibitors of human leukocyte clastase (HLE), cathepsin G (Cat G), and proteinase 3 (PR3), as well as antimycobacterium and central nervous system agents [9,12–14]. Recently, different saccharin-based antagonists have been recognized for their interferon-signaling pathways, showing antitumor activity through the inhibition of cancer-related isoforms in humans [15]. It should also be noted that the first non-benzoannelated 4-amino-2,3-dihydroisothiazole 1,1-dioxide, which lacks a 3-oxo group, has been described and shows anti-HIV-1 activity. Additionally, saccharin and isothiazolyl derivatives have been used in agriculture as herbicides, fungicides, and pesticides [16].

**Figure 1.** Structures of 1,2-benzisothiazole-3-one 1,1-dioxide (**1**, saccharin), 1*H*-tetrazole (**2**), and 1,3,4-thiadiazole (**3**).

Tetrazole (CN4H2) and its derivatives have attracted much attention as well due to their practical applications. The tetrazolic acid fragment –CN4H has acidity similar to the carboxylic acid group –CO2H and is almost allosteric with it, but it is metabolically more stable at the physiologic pH [17]. Hence, synthetic methodologies leading to the replacement of –CO2H groups by –CN4H groups in biologically active molecules are of major relevance [18]. Indeed, the number of patent claims and publications related to medicinal uses of tetrazolyl derivatives continues to grow rapidly and cover a wide range of applications: tetrazoles have been found, for instance, in compounds with antihypertensive, antiasthmatic, antitubercular, antimalarial, and antibiotic activity [19–22]. Several tetrazole derivatives have shown potential as anticonvulsants and anticancer and anti-HIV-1 drugs [23–25]. Tetrazoles have also had important applications in agriculture as plant growth regulators, herbicides, fungicides [26], and stabilizers in photography and photoimaging [27]. Due to the high enthalpy of formation, tetrazole decomposition results in the liberation of two nitrogen molecules and a significant amount of energy. Therefore, several tetrazole derivatives have been explored as explosives, propellant components for missiles, and gas generators for airbags (applicable to the automobile industry) [28]. In addition, various tetrazole-based compounds have good coordination properties and are able to form stable complexes with several metal ions. This ability is successfully used in analytical chemistry for the removal of heavy metal ions from liquids and in chemical systems formulated for metal protection against corrosion [29]. Many physical, chemical, physicochemical, and biological properties of tetrazoles are closely related to their ability to behave as acids and bases. In the tetrazole ring, the four nitrogen atoms connected

in succession are able to be involved in proteolytic processes. This heterocyclic system is unusual in structure and unique in terms of acid–base characteristics.

In line with what is mentioned above for saccharin and tetrazole derivatives, the 1,3,4-thiadiazole scaffold represents an important class of core structures that are of great interest mainly because of their various biological activities and respective therapeutic applications. The 1,3,4-thiadiazole ring is a very weak base, possesses relatively high aromaticity, and is moderately stable in aqueous acid solutions although it is vulnerable to ring cleavage with an aqueous base [30]. Besides, this heterocyclic ring is very electron-deficient due to the electron-withdrawing effect of the nitrogen atoms and is relatively inert toward electrophilic substitution but susceptible to nucleophilic attack, whereas when substitutions are introduced into the 2′ or 5′ position of this ring, it is highly activated and readily reacts to produce varied derivatives [31]. To some extent, these specific properties lead to the application of 1,3,4-thiadiazole derivatives in pharmaceutical, agricultural, and material chemistry. Therefore, several 1,3,4-thiadiazole-based compounds display a broad spectrum of biological activities, such as antimicrobial [32], antituberculosis [33], antioxidant [34], anti-inflammatory [35], anticonvulsant [36], antidepressant, anxiolytic [37], antihypertensive [38], anticancer [39], and antifungal activity [40]. The most prominent thiadiazole derivative is possibly the acetazolamide [*N*-(5-sulfamoyl-1,3,4-thiadiazol-2-yl)acetamide], a very well-known carbonic anhydrase inhibitor that is used in the treatment of glaucoma [41], high-altitude illness [42], epileptic seizures [43], idiopathic intracranial hypertension [44], hemiplegic migraine [45], cystinuria [46], obstructive sleep apnea [47], and congenital myasthenic syndromes [48].

The search for new molecular entities, whether natural or synthetic, with relevant pharmacological activities for effective applications in medical practices continues to be a hot topic in health science as well as in science as a whole. This permanent search is well documented in the immense scientific literature and is mainly supported by the fact that many pathogenic organisms are able to develop mechanisms of resistance to high-activity medicines in their early lives [49–54]. In this context, and given our prior interest in the synthesis, reactivity, and bioactivity of tetrazole and thiazole derivatives, our collaborative interest was piqued by the great potential of these heterocycles for medical applications. Herein, we report on the antimicrobial, antioxidant, and cytotoxic activities of four mixed-azole compounds of benzisothiazole–tetrazolyl and benzisothiazole–thiadiazolyl.

#### **2. Results and Discussion**

#### *2.1. Chemistry*

The 3-chloro-1,2-benzisothiazole 1,1-dioxide (**2**), one of the strategic building blocks for the synthesis of the studied molecules, was first prepared through the halogenation of saccharin (**1**), as previously described (Scheme 1 (A)) [55]. The three saccharin–tetrazolyl analogs (TS (**5**), 2MTS (**6**), and TSMT (**7**)) were synthetized through a combination of the amino-tetrazoles **3**, **4a**, and **4b** with **2**, following a nucleophilic substitution reaction of the chloride anion by the amine functionality (Scheme 1 (B)). Similarly, compound **9** (MTSB) was prepared by coupling **2** and 5-methyl-1,3,4-thiadiazole-2-thiol (**8**) (Scheme 1 (C)). All of the reactions proceeded smoothly under experimental protocols originally developed by us [56–59], affording crystalline products in reasonable to very good yields.

**Scheme 1.** Synthesis of saccharin–tetrazolyl (TS, 2MTS, and TSMT) and saccharin–thiadiazolyl (MTSB) derivatives.

#### *2.2. Biological Assays*

#### 2.2.1. Antimicrobial Activity

The assessment of the antibacterial properties of compounds TSMT, MTSB, TS, and 2MTS was determined through minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) values against Gram-positive (*Staphylococcus aureus* and *Enterococcus faecalis*), Gram-negative (*Pseudomonas aeruginosa* and *Escherichia coli*), and yeast (*Saccharomyces cerevisiae* and *Candida albicans*) strains obtained from the American Type Culture Collection (ATCC) (Table 1). The MIC value corresponding to the lowest concentration at which no visible growth was observed was assessed by the microdilution method [60]. For MBC evaluation, the bacterial suspension in the wells was homogenized, serially diluted, spread in triplicate on appropriate medium, and incubated at 37 ◦C. All compounds and respective positive controls were tested at the same concentration of 500 µg/mL.


**Table 1.** Minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) values of TSMT,MTSB, TS, and 2MTS (obtained through the microdilution method against Gram-positive bacteria, Gram-negative bacteria, and yeast strains (in µM)).

VAN: vancomycin; NOR: norfloxacin; NYS: nystatin; nt: not tested. Data are the median values of at least three replicates.

The results of the antimicrobial assay showed that the compounds tested were bacteriostatic. This was concluded because the MBC values were much higher than the MIC values (see Table 1). The compounds were more active against the Gram-negative *P. aeruginosa* and the Gram-positive *S. aureus* bacteria. However, it seemed that the MIC and MBC values were more similar against *E. coli*. Additionally, antimicrobial tests were performed at different pH values (pH 4.0, 7.0, and 9.0), also using the microdilution method (see Tables 2–4). The microorganisms used in these tests were chosen based on the initial results from the antimicrobial screening of Gram-positive bacteria and comprised four Gram-positive strains, namely *S. aureus* CIP6538, *E. faecalis* ATCC 29212, methicillin-resistant *S. aureus* CIP106760 (MRSA), and vancomycin-resistant *E. faecalis* ATCC51299 (VRE).

**Table 2.** MIC and MBC values of TSMT, MTSB, TS, and 2MTS (obtained through the microdilution method against Gram-positive bacteria at pH 4.0).


VAN: vancomycin; NOR: norfloxacin; NYS: nystatin; nt: not tested. Data are the median values of at least three replicates.

**Table 3.** MIC and MBC values of TSMT, MTSB, TS, and 2MTS (obtained through the microdilution method against Gram-positive bacteria at pH 7.0).


VAN: vancomycin; NOR: norfloxacin; NYS: nystatin; nt: not tested. Data are the median values of at least three replicates.


**Table 4.** MIC and MBC values of TSMT, MTSB, TS, and 2MTS (obtained through the microdilution method against Gram-positive bacteria at pH 9.0).

VAN: vancomycin; NOR: norfloxacin; NYS: nystatin; nt: not tested. Data are the median values of at least three replicates.

The results of the antimicrobial assays at the different pH values showed that lowering the medium pH to 4.0 had a positive effect on the compounds MTSB, TS, and 2MTS against the methicillin-resistant *S. aureus* CIP106760 (MRSA). Overall, it was attested that MTSB at pH 4.0 was the most active derivative against all Gram-positive strains. It should be noted that MTSB was the sole compound comprising the thiadiazole function, and presumably, the different activity must be correlated with the type of heterocycle ring. The results for pH 7.0 and 9.0 did not provide better results than those previously obtained.

In terms of pH-dependent antimicrobial mechanisms, it should be emphasized that several antimicrobial peptides (AMPs) have increasingly been reported as potent antibiotics that utilize pH-dependent antimicrobial mechanisms [61]. Some of these antibiotics display high pH optima related to their antimicrobial activity and show activity against microbes that present low pH optima, which reflects the acidic pH generally found at their sites of action, namely the skin. This effect should be comparable to our compounds and could be the explanation for the high antimicrobial activity of our compounds at low pH. Several pH-dependent AMPs and other antimicrobial proteins have been developed for medical purposes and have successfully gone through clinical trials, namely kappacins, LL-37, histatins, lactoferrin, and their derivatives. The major examples of the therapeutic applications of these antimicrobial compounds include wound healing as well as the treatment of multiple infections. Generally, such applications involve topical administration, a source of novel biologically active agents that could aid in the fulfilment of the urgent need for alternatives to conventional antibiotics, helping to avert a return to the pre-antibiotic era: our compounds could be a key development.

#### 2.2.2. Antioxidant Activity

The antioxidant activity of the studied compounds was evaluated using a DPPH assay, which evaluates the potential of test samples to quench DPPH radicals via hydrogen-donating ability. The antioxidant agents convert DPPH into a stable diamagnetic molecule, 1-1diphenyl-2-picryl hydrazine, through electron or hydrogen transfers. A color change from purple to yellow indicates the increasing radical scavenging activity of the test compounds. Herein, it was observed that TSMT and MTSB derivatives possessed a high free radical scavenging ability (see Figure 2). These two molecules had the highest reducing power, indicating that they were good electron/hydrogen donors and could prevent oxidative stress.

**Figure 2.** Antioxidant activity tested at a concentration of 10 µg/mL. The mean value ± SD was calculated from three independent experiments and compared to butylated hydroxytoluene (BHT) (∗ *p* < 0.05; ∗∗ *p* < 0.001).

#### 2.2.3. Brine Shrimp Lethality Bioassay (General Toxicity)

The *Artemia salina* test is a known, simple, fast, and low-cost test and was used in this investigation to check the general toxicity of the compounds. As a general rule, it was observed that all of the compounds exhibited low toxicity in the *Artemia salina* model (Figure 3). Nevertheless, it should be emphasized that the MTSB and TSMT derivatives presented with higher toxicity, making them potential lead therapeutic agents, and thus they must further be tested using different cell- and microorganism-based assays. \*\*

**Figure 3.** General toxicity screening at a concentration of 10 ppm using the *Artemia salina* test. The mean value ± SD was calculated from three independent experiments and compared to salt (∗∗ *p* < 0.001).

2.2.4. Cell Viability after Treatment with MTSB, TS, TSMS, and 2MTS in H7PX Glioma Cells (IV Grade)

In the course of this study, H7PX cells were treated with a concentration range of 0–100 µg/mL of the four compounds (MTSB, TS, TSMS, and 2MTS) over 24 h, after which the percentage of cell viability was determined by an MTT assay. It was demonstrated that none of the tested compounds

reduced the viability of human H7PX cells in the stated range of concentration (0–100 µg/mL) after 24 h (Figure 4). None of the compounds' IC<sup>50</sup> values were reached. Similar effects were observed with 48 h of incubation (data not shown).

**Figure 4.** Effect of MTSB, TS, TSMT, and 2MTS treatment on the viability of H7PX glioma cells after 24 h of incubation (data are reported as means ± SD of three determinations).

#### **3. Experimental Section**

#### *3.1. Chemistry*

#### 3.1.1. General

Unless indicated otherwise, solvents and starting materials were obtained from Sigma. All chemicals used were of reagent grade without further purification before use. Column chromatography was performed using silica gel 60 MN, and aluminum-backed silica gel Merck 60 F254 plates were used for analytical thin-layer chromatography (TLC). Melting points were recorded and are uncorrected. <sup>1</sup>H and <sup>13</sup>C NMR spectra were recorded at room temperature on a Bruker Avance II 400 (UltraShield™ Magnet, Billerica, MA, USA) spectrometer operating at 400 MHz (1H) and 101 MHz (13C). The chemical shifts are reported in ppm using TMS (tetramethylsilane) as an internal standard. Carbon, hydrogen, and nitrogen elemental analyses were carried out by the Microanalytical Service of the Instituto Superior Técnico—University of Lisbon. FT-IR spectra (4000–400 cm–1) were recorded on a VERTEX 70 (Bruker, Billerica, MA, USA) spectrometer using KBr pellets. Mass spectra were obtained on a VG 7070E mass spectrometer through electron ionization (EI) at 70 eV.

#### 3.1.2. Synthetic Protocols

The synthesis of 3-chloro-1,2-benzisothiazole-1,1-dioxide (**2**), 2-methyl-(2*H*)-tetrazole-5-amine (**4a**), 1-methyl-(1*H*)-tetrazole-5-amine (**4b**), *N*-(1*H*-tetrazol-5-yl)-*N*-(1,1-dioxo-1,2-benzisothiazol-3-yl) amine (**5**) (TS), *N*-(2-methyl-2*H*-tetrazol-5-yl)-*N*-(1,1-dioxo-1,2-benzisothiazol-3-yl) amine (**6**) (2MTS), *N*-(1-methyl-2*H*-tetrazol-5-yl)-*N*-(1,1-dioxo-1,2-benzisothiazol-3-yl) amine (**7**) (TSMT), and 3-[(5-methyl-1,3,4-thiadiazol-2-yl)sulfanyl]-1,2-benzisothiazole 1,1-dioxide (**9**) (MTSB) was carried out as previously described [55,56,58,59].

#### *3.2. Biologic Activities*

#### 3.2.1. Antioxidant Activity (DPPH Method)

The antioxidant activity of all the compounds was measured by the DPPH method, as described by Rijo et al. [62]. Accordingly, a mixture containing 10 µL of sample and 990 µL of DPPH solution (0.002% in methanol) was incubated for 30 min at room temperature followed by absorbance measurements at 517 nm against the corresponding blank sample. The antioxidant activity of each compound was calculated using Equation (1). *AA* denotes the antioxidant activity, *ADPPH* is the absorption of DPPH against the blank, and *Asample* represents the absorption of the compound or control against the blank. All tests were carried out in triplicate at a sample concentration of 10 µg/mL. The reference standard used for this procedure was butylated hydroxytoluene (BHT) in the same conditions as the samples:

$$\text{AA} = \frac{\text{A}\_{\text{DPPH}} - \text{A}\_{\text{Sample}}}{\text{A}\_{\text{DPPH}}} \times 100\% \tag{1}$$

#### 3.2.2. Brine Shrimp Lethality Bioassay (General Toxicity)

The general toxicity of the compounds was evaluated by the use of a test of lethality to *Artemia salina* (brine shrimp) [63]. Concentrations of 10 ppm of each sample were tested. The number of dead larvae was recorded after 24 h and was used to calculate the lethal concentration (%) according to Equation (2) (where *TotalA. salina* = the total number of larvae in the assay and *AliveA. salina* = the number of alive *A. salina* larvae in the assay):

$$\text{Total concentration} = \frac{\text{Total}\_{\text{A. salina}} - \text{Alive}\_{\text{A. salina}}}{\text{Total}\_{\text{A. salina}}} \times 100\% \tag{2}$$

#### 3.2.3. Cell Culture

An H7PX primary glioblastoma cell line was cultivated in DMEM (Biowest, Nuaollé, France) supplemented with 10% FBS (Euroclone, Pero, Italy), 100 IU/mL penicillin (Sigma-Aldrich, Saint Louis, MO, USA), 100 µg/mL streptomycin (Sigma-Aldrich, Saint Louis, MO, USA) and 50 µg/mL gentamicin (Biowest, Nuaollé, France) in a humidified atmosphere (5% CO2, 37 ◦C).

#### 3.2.4. In Vitro Cell Viability by MTT Assay

An MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide) assay was employed to measure the viability of H7PX cells (glioma cells in IV grade derived from patient) treated with different concentrations of TS (**5**), 2MTS (**6**), TSMT (**7**), or MTSB (**9**). Cells were seeded at 1 × 10<sup>4</sup> cells per well in 96-well culture plates and were left overnight before treatments for attachment. Subsequently, the cells were incubated for 24 h with all compounds over a range of concentrations: 0.0 (control), 0.39, 0.78, 1.56, 3.13, 6.25, 12.5, 25, 50, and 100 µg/mL. Following this, the cells were incubated with 0.5 mg/mL of MTT at 37 ◦C for 1.5 h. After that, the MTT was carefully removed, and DMSO (100 µL) was added to each well and vortexed at low speed for 5 min to fully dissolve the formazan crystals. Absorbance was measured at 570 nm with a reference at 630 nm using a Bio-Tek Synergy HT Microplate Reader (Bio-Tek Instruments, Winooski, VT, USA). All experiments were repeated in triplicate. Cell viability was expressed as a percentage relative to the untreated cells, which was defined as 100%. The study was approved by the Ethical Commission of the Medical University of Lodz, and informed consent was obtained from the patients (Nr. RNN/194/12/KE).

#### 3.2.5. Antimicrobial Activity

Microorganisms and growth conditions: The strains used in this study comprised *Staphylococcus aureus* (ATCC 25,923 and CIP 106760), *Enterococcus faecalis* (ATCC 51,299 and ATCC29212), *Escherichia coli* ATCC 25922, *Pseudomonas aeruginosa* ATCC 27,853, and the yeasts *Candida albicans* ATCC 10,231 and *Saccharomyces cerevisiae* ATCC 2601. All bacteria were grown at 37 ◦C in Mueller–Hinton broth (Biokar Diagnostics, Beauvais, France), and the yeasts were grown in Sabouraud dextrose agar (Biokar Diagnostics, Allone, France).

Microdilution method (MIC determination): The minimum inhibitory concentrations (MICs) of the compounds (dissolved in the respective pH aqueous solutions) were evaluated using a twofold serial broth microdilution assay (CLSI, 2011) in Mueller–Hinton broth (MHB, Biokar Diagnostics, Beauvais, France). Overnight cultures were diluted in MHB with increasing concentrations of each compound (in µM). Vancomycin (VAN), norfloxacin (NOR), and nystatin (NYS) were used as positive controls for Gram-positive bacteria, Gram-negative bacteria, and yeasts, respectively. The negative control for the aqueous solutions at different pH values showed no inhibition growth. The cultures were incubated for 24 h at 37 ◦C, and Optical Density at 620 nm was measured using a Microplate Reader (Thermo Scientific Multiskan FC, Loughborough, UK). Assays were carried out in triplicate for each tested microorganism.

Minimum bactericidal concentration (MBC) assessment: To define the minimum bactericidal concentration (MBC) for each set of wells in the MIC determination, a loopful of agar was collected from the wells without any growth and inoculated on sterile Mueller–Hilton medium broth (for bacteria) through streaking. Plates inoculated with bacteria were incubated at 37 ◦C for 24 h. After incubation, the lowest concentration was noted as the MBC (for bacteria) at which no visible growth was observed.

#### 3.2.6. Statistical Analysis

The values in this study are expressed as means ± SD. The Shapiro–Wilk test was used for verification of the normality of the data. Statistical differences were determined by one-way ANOVA. The results were analyzed using STATISTICA 12.0 software (StatSoft, Tulsa, OK, USA). Differences of *p* < 0.05 were considered statistically significant.

#### **4. Conclusions**

The antimicrobial, antioxidant, and cytotoxic activities of three saccharin–tetrazolyl (TS, TSMT, and 2MTS) derivatives and one saccharin–thiadiazolyl (MTSB) derivative were addressed throughout this investigation. The antimicrobial activity of the synthesized compounds was evaluated against a series of Gram-positive and Gram-negative bacteria and yeast strains. An evaluation of the MIC and MBC values of the four derivatives was completed at pH 4.0, 7.0, and 9.0 against four Gram-positive strains (*S. aureus*, *E. faecalis*, *S. aureus* (MRSA), and *E. faecalis* (VRE)), showing high values for the MIC and MBC at pH 4.0 (ranging from 3.42 to 473.0 µM). It was attested that the derivative MTSB, the sole compound comprising the thiadiazole function, was the most active against all of the considered Gram-positive strains at pH 4.0.

In addition, the antioxidant activity of the compounds (calculated by using the DPPH method) was the highest value for TSMT (90.29% compared to the BHT positive control of 61.96%). Finally, we demonstrated for the first time that the TS, TSMT, 2MTS, and MTSB compounds did not show in vitro cytotoxic effects on H7PX glioma cells.

The present study exposed the influence of the pH of the medium on the antimicrobial activity of the TS, TSMT, 2MTS, and MTSB compounds, which is similar to well-described antimicrobial peptide antibiotics. Therefore, the use of these kinds of molecules to produce new antibiotics should be considered in the future, although further studies are needed to confirm this hypothesis.

**Author Contributions:** L.M.T.F. and P.R. designed the study, analyzed the data, and wrote the paper; E.N., P.S., J.M.A., M.T., and T.S. performed the biological assays; L.M.T.F. and L.C. performed the synthesis and ´ characterization of the compounds; M.L.S.C. and A.J.L.P. were responsible for supervision and reviewing the draft.

**Funding:** This work was partially supported by the Foundation for Science and Technology (FCT), Portugal ((UID/QUI/00100/2019) and (UID/MULTI/04326/2019 – CCMAR)). L.M.T.F. expresses gratitude to FCT for the post doc fellowship (SFRH/BPD/99851/2014) and work contract n◦ IST-ID/115/2018.

**Conflicts of Interest:** The authors declare no conflicts of interest.

### **References**


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