*Article* **Manipulating Estrogenic/Anti-Estrogenic Activity of Triphenylethylenes towards Development of Novel Anti-Neoplastic SERMs**

**Heba E. Elnakib 1,† , Marian M. Ramsis 1,† , Nouran O. Albably 1,† , Merna A. Vector 1 , Jan J. Weigand 2 , Kai Schwedtmann 2 , Jannette Wober 3, \*, Oliver Zierau 3 , Günter Vollmer 3 , Ashraf H. Abadi 1 and Nermin S. Ahmed 1, \***


**Abstract:** Selective estrogen receptor modulators (SERMs) act as estrogen receptor (ERα) agonists or antagonists depending on the target issue. Tamoxifen (TAM) (a non-steroidal triphenylethylene derivative) was the first SERM approved as anti-estrogen for the treatment of metastatic breast cancer. On the hunt for novel SERMs with potential growth inhibitory activity on breast cancer cell lines yet no potential to induce endometrial carcinoma, we designed and synthesized 28 novel TAM analogs. The novel analogs bear a triphenylethylene scaffold. Modifications on rings **A**, **B,** and **C** aim to attenuate estrogenic/anti-estrogenic activities of the novel compounds so they can potentially inhibit breast cancer and provide positive, beneficial estrogenic effects on other tissues with no risk of developing endometrial hyperplasia. Compound **12** (*E/Z*-1-(2-{4-[1-(4-Chloro-phenyl)-2-(4-methoxyphenyl)-propenyl]-phenoxy}-ethyl)-piperidine) showed an appreciable relative ERα agonistic activity in a yeast estrogen screen (YES) assay. It successfully inhibited the growth of the MCF-7 cell line with GI<sup>50</sup> = 0.6 µM, and it was approximately three times more potent than TAM. It showed no potential estrogenicity on Ishikawa endometrial adenocarcinoma cell line via assaying alkaline phosphatase (AlkP) activity. Compound **12** was tested in vivo to assess its estrogenic properties in an uterotrophic assay in an ovariectomized rat model. Compared to TAM, it induced less increase in wet uterine wet weight and showed no uterotrophic effect. Compound **12** is a promising candidate for further development due to its inhibition activity on MCF-7 proliferation with moderate AlkP activity and no potential uterotrophic effects. The in vitro estrogenic activity encourages further investigations toward potential beneficial properties in cardiovascular, bone, and brain tissues.

**Keywords:** tamoxifen; CYP2D6; MCF-7; Ishikawa cells; SERM; TNBC; uterotrophic

#### **1. Introduction**

Selective estrogen receptor modulator (SERM) refers to a structurally diverse group of compounds that binds to both estrogen receptor subtypes ERα and/or ERβ despite lacking the estrogen steroid moiety. Whereas estrogens typically exert ER agonist effects, SERMs confer mixed functional ER agonist or antagonist activity depending on the target tissue [1]. An ideal SERM would have ER agonist activity in tissues where mimicking the action of estrogens is desirable (e.g., skeletal, cardiovascular, and central nervous systems), and lack of estrogenicity in tissues where estrogens have been shown to induce cancer initiation

**Citation:** Elnakib, H.E.; Ramsis, M.M.; Albably, N.O.; Vector, M.A.; Weigand, J.J.; Schwedtmann, K.; Wober, J.; Zierau, O.; Vollmer, G.; Abadi, A.H.; et al. Manipulating Estrogenic/Anti-Estrogenic Activity of Triphenylethylenes towards Development of Novel Anti-Neoplastic SERMs. *Int. J. Mol. Sci.* **2021**, *22*, 12575. https://doi.org/ 10.3390/ijms222212575

Academic Editor: Angela Stefanachi

Received: 10 October 2021 Accepted: 16 November 2021 Published: 22 November 2021

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**Copyright:** © 2021 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 (https:// creativecommons.org/licenses/by/ 4.0/).

and growth (e.g., breast and endometrium) [2]. This definition led to investigations on the clinical profile of an ideal SERM. An ideal SERM prevents bone loss and fractures yet does not stimulate endometrial hyperplasia. It also provides relief of hot flushes and other menopausal symptoms. It should not increase the risk of coronary heart disease, stroke, or deep vein thrombosis. The first-generation triphenylethylene SERM included tamoxifen (TAM) and toremifene. Both SERMs are far from being ideal [3].

TAM (I) (a non-steroidal triphenylethylene derivative) was the first SERM approved as anti-estrogen for the treatment of metastatic breast cancer. It is now widely used as adjuvant chemotherapy for the treatment of hormone-dependent metastatic breast carcinoma in postmenopausal women. Although TAM (I) has been very successful in treating breast cancer, some side effects such as thromboembolic events, vasomotor symptoms, and an increased risk of endometrial hyperplasia are associated with TAM treatment [4].

TAM (I) is regarded as a prodrug that is metabolized to the more active metabolites: 4-OH-TAM (II) and endoxifen (III) Figure 1 [5]. Compared to the parent drug, those metabolites have 100-times more affinity to the ER. This metabolism is mainly mediated via cytochrome P450 (CYP) enzymes, specifically the CYP2D6 and CYP3A4 isoforms. Pharmacogenetics revealed the polymorphic nature of the CYP2D6 enzyme. CYP2D6 poor metabolizers (based on *CYP2D6*\*4 and \*6) were reported to benefit less from TAM compared with extensive metabolizers [6–8].

**Figure 1.** Structures of TAM (**I**), 4-OH-TAM (**II**), and endoxifen (**III**).

The different phenotypes lead to different plasma concentrations of active metabolites among patients of different populations, and hence different clinical outcomes and may lead to drug resistance. Thus, to overcome TAM resistance, TAM is perceived as a clinical target in oncology personalized medicine [9–11].

On the hunt for novel SERMs that possess potential growth inhibitory activity on breast cancer cell lines yet lack the potential to induce endometrial carcinoma, we designed and synthesized 28 novel TAM analogs. The novel analogs bear a triphenylethylene scaffold. Modifications on rings **A**, **B**, and **C** aim to attenuate estrogenic/anti-estrogenic activities of the novel compounds so they can potentially inhibit breast cancer and provide positive estrogenic effects on bones and cardiovascular system without affecting endometrial tissues.

Structural modifications included introducing a chlorine atom at position 4 on ring **C** in all analogs; this ensures the blockage of the site of *para*-hydroxylation; thus, those analogs can bypass *para*-hydroxylation by polymorphic CYP2D6. The effect of this modification on the compounds estrogenic/anti-estrogenic properties is investigated.

The introduction of fluorine into a molecule can productively influence conformation, p*K*a, intrinsic potency, membrane permeability, metabolic pathways, and pharmacokinetic properties [12]. Based on these findings, ring **A** was kept unsubstituted or modified to 4-methoxy phenyl, 4-methoxy-3-fluoro phenyl, or 4-fluoro-3-methoxy phenyl. The introduction of a small, highly electronegative fluorine atom on ring **A** can affect the novel analogs stability and lipophilicity. The fluorine atom can further affect the binding affinity either directly or by affecting the polarity of the adjacent methoxy groups.

Previous literature focused on the effect of substitution on position 4 of ring **B** [13]; additionally, recent studies even suggested different substituents on ring **B** to design a homodimeric ER ligand that can act as ER antagonist and SERD (selective estrogen receptor

degrader) [14]. In our work, the effect of the length of the alkoxy chain, size, and bulkiness of *N*-substituents and cyclization are thoroughly studied. The novel compounds were depicted in Table 1.

**Table 1.** Synthetic scheme: Preparation of compounds **1**–**28**.


All synthesized compounds were tested for their relative activity in β-galactosidase in a yeast estrogen screen (YES) assay. Compounds were tested for their relative estrogenic/ anti-estrogenic activities in comparison to positive and negative controls, respectively. YES assay is a gene reporter assay where the DNA sequence of human ERα is integrated into the yeast genome completed with an expression plasmid carrying estrogen response elements (ERE) in the promoter controlling the expression of the reporter gene *lacZ* (encoding the enzyme β-galactosidase). In the presence of estrogenic compounds, β-galactosidase is

synthesized and secreted into the medium, where it converts the chromogenic substrate chlorophenol red-β-D-galactopyranoside (CRPG) from a yellow to a red product, whose absorbance is measured. Agonistic activity is measured directly, whereas antagonistic activity is measured in terms of reduction in color formation in the presence of 0.5 nM/1 nM estradiol (E2) [15–17]. Despite the ability of the YES assay to differentiate between agonists and antagonists, it becomes more and more apparent that compounds exhibit an organselective mode of action [18]. Therefore, we decided to test the novel compounds in an organ-specific in vitro model using the human Ishikawa endometrial adenocarcinoma cell line [19,20].

Alkaline phosphatase (AlkP) activity in these human endometrial cancer cells is markedly stimulated by estrogens [9]. In addition and in contrast to yeast assays, which do not mimic human metabolism, the Ishikawa cells, such as normal uterine cells, possess the important capacity to metabolize the compounds, which reflects their true estrogenic activity [21,22].

The anti-proliferative effects of the novel analogs were tested in vitro by the National Cancer Institute (NCI) on a panel of 60 human tumor cell lines at 10 µM. Compounds that elicited mean growth inhibition ≥50% were selected by the NCI for 5-dose testing. The concentration for 50% of maximal inhibition (GI50), total growth inhibition (TGI), and half-maximal lethal concentration (LC50) was measured for each cell line.

Compounds showing appreciable estrogenic activity in the YES assay and that were able to inhibit the growth of MCF-7 cancer cell lines yet with low estrogenic activity on Ishikawa endometrial adenocarcinoma cells might serve as potential ideal SERMS. Compounds **12** and **19** were therefore selected for the in vivo experiments to assess their estrogenic properties in an uterotrophic assay in an ovariectomized rat model. The in vivo uterotrophic rat assay is the gold standard assay to test for the estrogenic effect of compounds; the assay uses adult ovariectomized (OVX) female rats where there is no significant source of endogenous estrogens. Compounds that have estrogenic effects cause uterotrophic response due to the imbibition of water and growth of the uterine cells. Statistically significant uterine weight increases compared to controls provide a positive result [23–29]. Adopting both in vivo and in vitro assays was inevitable due to the limitations of each assay. The cell lines are not properly able to recapitulate the in vivo environment of the uterus within the body. On the other hand, the rat uterotrophic assay merely considers the uterine weight gain as an endpoint of estrogenicity without taking into account all factors that play a role in exerting an estrogenic effect on the organ and body [30].

All our compounds were biologically assayed as *E-Z* mixtures due to synthetic challenges and failure in separating the isomers using available chromatographic techniques. We adopted an in silico model to postulate the isomer with the lowest binding energy. The model also investigates the full agonistic activity of compound **3** despite the lack of an OH group on ring **C**. This group was reported to be essential for ER binding affinity of most synthetic ER ligands.

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

#### *2.1. Chemistry Discussion*

Compounds (**1**–**4**) were synthesized using standard McMurry coupling reaction of 4-Chloro-4-hydroxybenzophenone with commercially available ketones using titanium tetrachloride/zinc as a catalyst to yield four condensation products. The condensation products (**1**–**4**) were then treated with the appropriate base hydrochloride salts in dimethyl formamide (DMF) in the presence of potassium carbonate to form ethers (**5**–**28**) [31]. The formation of all compounds and their purity were confirmed via UPLC-ESI MS. All compounds were obtained as a mixture of E-Z isomers, as shown from UPLC-UV chromatograms. Some chromatograms showed distinct two peaks of nearly similar area (1:1.1) and having the same molecular ion peak (M+H)<sup>+</sup> . Attempts to isolate the E-Z isomers using column chromatography as well as preparative HPLC were not successful. <sup>1</sup>H-NMR showed peaks integrating for double the number of protons, further confirming

the formation of a mixture of E-Z isomers. <sup>13</sup>C-NMR further confirmed the formation of isomers since most of the signals were duplicated. Such duplication of signals has been previously reported by Bedford and Richardson [32]. Their masses were confirmed via their molecular ion peaks [M+H]<sup>+</sup> and [M+H+2]<sup>+</sup> due to the presence of chlorine atoms in all compounds. As previously employed in similar work in the literature, compounds were assayed biologically as E-Z mixtures [33–36].

#### *2.2. Anti-Estrogenic Assays*

All compounds lacked significant anti-estrogenic on the ERα except compounds **27** and **28,** which were slightly able to antagonize the *β*-galactosidase reporter gene activity induced by 1 nM E2 by 11% and 12%, respectively. It seems that the para chlorine substitution at ring **C** has a detrimental effect on the anti-estrogenic activity. This modification has blocked the action of CYP2D6 and therefore prevented the formation of the anti-estrogenic hydroxy metabolite. It is reported that 4-OH-TAM and endoxifen, the active metabolites of TAM, have higher anti-estrogenic potency than the parent drug, TAM [33]. The OH group at position 4 of 4-OH-TAM is presumed to be responsible for its higher anti-estrogenic activity compared to TAM. Additionally, studies have reported that the anti-estrogenic property of SERMs depends on the ability of the cationic nitrogen on the alkylaminoethoxy side chain on ring **B** to neutralize the charge of Asp 351 [37]. Our results showed that the presence of a basic alkylaminoalkoxy group without a phenolic OH on ring **C** or a phenyl ring prone to metabolic hydroxylation could not elicit anti-estrogenic activity regardless of the size and basicity of this group, as shown in compounds **5**–**26**. Having no tertiary amino group on ring **B** as shown in compounds **3** and **4** or blocking position 4 on ring **C** as shown in compounds **5**–**26** will mostly abolish the anti-estrogenic action and shift it toward estrogenic activity (Table 2).


**Table 2.** Relative β-galactosidase activity using YES assay (antagonistic activity).

\* Relative anti-estrogenic activity is compared to 0.5 nM/1 nM E2 (set as 1), compounds screened at a dose of 1 µM in presence of 0.5 nM/1 nM E2, respectively; compounds were screened in triplicates; \*\* n.d. = not determined. Compounds were not selected for anti-estrogenic assays due to their high estrogenic activity.

This drives us to the hypothesis that the alkylaminoethoxy side chain on ring **B** is not the only crucial factor for anti-estrogenicity. There are essentially two important features responsible for anti-estrogenic activity. A phenolic OH group is required for high-affinity binding to ER-forming crucial interactions (H-bonds) with Glu 353 and Arg 394 amino acids in the ligand-binding domain (LBD), and the alkylaminoalkoxy bulky group at ring **B** is essential for the ER antagonistic action where it forms a cationic interaction with Asp 351 amino acid of the ER [38].

#### *2.3. Estrogenic Assays*

All synthesized compounds were tested for their relative β-galactosidase activity in a yeast estrogen screen (YES) assay at a concentration of 1 µM using DMSO as control (set as 1). The hydroxylated analogs **3** and **4** showed EC<sup>50</sup> values of 40.1 nM and 258 nM, respectively. E2, the endogenous ligand, showed an EC<sup>50</sup> = 0.528 nM. The remarkable potency of the two novel analogs can be attributed to the introduction of a chloro group at the para position of ring **C**, the hydroxyl group of ring **B,** and the nature of the substituents on ring **A**.

Compound **3** (EC<sup>50</sup> = 40 nM) bears a methoxy group at position 4 and a fluoro group at position 3 on ring **A**, and compound **3** showed six-fold more potency than its positional isomer compound **4** (EC<sup>50</sup> = 258 nM). It seems that a methoxy group at position 4 is essential for agonistic activity.

This could further support the hypothesis that the introduction of a chloro group at ring **C** resulted in an estrogenic property, and the presence of an OH group at ring **B** allows better fitting into the receptor, ensures higher binding affinity, and locking the receptor drug complex into an agonistic conformation.

Replacing the OH group with different alkylaminoalkoxy side chains did not abolish the estrogenic action yet caused a decrease in activity. Comparing compounds (**5**–**9**) bearing a chloro group at ring **C**, unsubstituted ring **A** but different alkylaminoalkoxy side chains, compound **9** with an azepanethoxy side chain at ring **B** induced high relative β-galactosidase activity of 6.74 compared to control; a bulky cyclized side chain on ring **B** seems to improve estrogenic activity.

Compounds (**10**–**14**) bear a methoxy substituent on ring **A**. Both compounds **10** and **13** were the most potent congeners. They bear a dimethylaminopropoxy side chain and a morpholinylethoxy side chain, respectively, on ring **B** (relative β-galactosidase activity = 11.61 and 12.41, respectively). The para methoxy substituent led to an increase in relative estrogenic activity for compounds **10** and **13** compared to their congeners **5** and **8**. A remarkable decrease in relative estrogenic activity was observed for compound **14** compared to its congeners **9**; this may be explained by the fact that the bulky azepanylethoxy group displaced the methoxy substituent of ring **A** outside the binding pocket leading to a possible steric clash.

Compounds (**15**–**21**) bear 3-fluoro 4-methoxy on ring **A,** whereas compounds (**21**–**28**) bear 3-methoxy 4-fluoro substituents on ring **A**. The alkylaminoethoxy side chains on ring **B** were extended to include dimethylaminoethoxy and diethylaminoethoxy side chains. For all compounds (**15**–**21**), the addition of a fluoro group at position **3** enhances the relative estrogenic activity compared to their structural isomers (**22**–**28**) except for compound **18**. The unexpected behavior of compound **18** may be attributed to the less lipophilic character of this compound and lower pK<sup>a</sup> value as a result of the morpholinylethoxy substituent on ring **B**. Compounds **15** and **17**, bearing a dimethylaminopropoxy side chain and a piperidinylethoxy side chain, respectively, showed relative estrogenic activities of 7.77 and 7.28, respectively. Compound **17** was the most potent among their series EC<sup>50</sup> = 252 ± 8 nM. Comparing compound **17** with compound **12**, compound **17** was two-fold more estrogenic at 1 µM, the introduction of a fluoro group at the meta position had a positive impact on estrogenic activity. Compound **19** bearing azepanylethoxy group on ring **B** showed relative estrogenic activities of 3.22 and EC<sup>50</sup> = 407 ± 86 nM, indicating that estrogenic activity is retained with bulky substituents. Compounds (**22**–**28**) were nearly equipotent. Modifying ring **A** to 3-methoxy 4-fluoro phenyl has resulted in a remarkable decrease in estrogenic activity. It seems that the methoxy substituent at the para position and fluoro substituent at the meta position of ring **A** is the main determinant factors for the higher agonistic action rather than the size or cyclization of substituents on ring **B** (Tables 3 and 4).


**Table 3.** Relative β-galactosidase activity using YES assay (agonistic activity).

\* Estrogenic activity is compared to DMSO (set as 1), compounds screened at a dose of 1 µM; compounds were screened in triplicates.

**Table 4.** EC<sup>50</sup> values (agonistic activity) of selected compounds.


\* n.c. = not calculable because no upper plateau is detectable; compounds were screened in triplicates.

#### *2.4. NCI Growth Inhibition Assays*

Compounds were submitted to the Developmental Therapeutics Program (DTP) of the National Cancer Institute (NCI). The program uses a panel of 60 human tumor cell lines representing nine tissue types, including leukemia, non-small cell lung cancer (NSCLC), melanoma, colon cancer, ovarian cancer, CNS cancer, renal cancer, prostate cancer, and breast cancer, to screen for potential new anti-cancer agents. SRB (sulforhodamine B) assay is the preferred high-throughput assay of the National Cancer Institute (NCI) and is the assay used in the NCI's lead compound screening program. Primary screening of synthesized compounds was performed by testing a single high dose of 10 µM in the full NCI-60 panel. The percent growth of treated cells relative to the no-drug control and relative to the time zero number of cells was measured, and a mean graph was provided. The percentage inhibition was then calculated by subtracting the values obtained from 100. In general, all compounds bearing an OH group (**3** and **4**) or a morpholinylethoxy side chain on ring **B** (**8**, **13**, **18,** and **25**) lacked anti-proliferative activity. They had the least percent mean growth inhibition and the lowest percent inhibition on human breast cancer MCF-7 cells. This may be attributed to the partial hydrophilicity of ring **B** (Table 5).


**Table 5.** Percent mean growth inhibition on 60 NCI tumor cell lines and on MCF-7 cells.

\* Data obtained from NCI in vitro disease-oriented human tumor cell screen (for details, see the work of [39]), compounds tested at a concentration of 10 µM in triplicates.

> Six compounds: **11** (67.76%), **12** (55.21%), **16** (77.24%), **17** (69.12%), **19** (60.79%), **28** (92.33%), showed mean percentage inhibition on all 60 cell lines higher than 50% and were escalated to a dose-dependency assay using five doses on the 60 cell panel. Five of the six compounds share two common features; they bear a para methoxy substituent on ring **A** and bear a cyclic alkylaminoethoxy group on ring **B**.

> In the dose-dependency assay, compounds were evaluated against the 60-cell panel at the five doses; 10−<sup>4</sup> M, 10−<sup>5</sup> M, 10−<sup>6</sup> M, 10−<sup>7</sup> M, and 10−<sup>8</sup> M. Dose-response curves for each cell line was drawn, and three response parameters are extracted by linear interpolation (GI50, TGI, LC50).

> To investigate SERM-like properties of compounds, looking at results from ER-positive cell lines is particularly important. The two most potent compounds on Erα-positive MCF-7 breast cancer cell line were compounds **11** (GI<sup>50</sup> = 0.89 µM) and **12** (GI<sup>50</sup> = 0.60 µM). They are almost twice as active as TAM (GI<sup>50</sup> = 1.58 µM; see Supplementary Materials). Both compounds bear a para methoxy substitution on ring **A** and a cyclic aminoethoxy group on ring **B,** namely a pyrolidine and piperidine, consecutively. The incorporation of the basic nitrogen in a cyclic structure enhances its basicity and significantly improves the anti-proliferative effect of the compounds.

> Compounds **16**, **17,** and **19** showed GI<sup>50</sup> = 2.41, 3.34, and 3.59 µM, respectively. Those compounds bear a para methoxy substituent and a meta fluorine substituent on ring **A**. They exhibited lower anti-proliferative activity on the MCF-7 breast cancer cell line compared to their congeners that lack a fluorine group on meta position, e.g., compounds **11**, **12**, and **14**. This suggests that the presence of a methoxy group increased the electron density on ring **A** resulting in a better anti-proliferative activity, whereas the introduction of an electronwithdrawing group such as fluorine at the meta position lowered the activity. We presumed that introduction of fluorine will increase compounds lipophilicity and therefore improve compounds' cellular uptake and growth inhibition potential. Switching the positions of the methoxy and fluorine substituents in compounds (**22**–**28**) deteriorated the antiproliferative activity except in compound **28** (GI<sup>50</sup> = 2.17 µM). This further confirms that fluorine develops essential interactions with specific targets involved in novel compounds' cytotoxic activities.

> It is worth mentioning that all six escalated compounds showed more potent antiproliferative activity than TAM on triple-negative breast cancer (TNBC) cell lines MDA-MDB-231/ATCC and BT-549. Compounds **17**, **19,** and **28** were more potent than TAM on Hs578T, whereas only compound **28** was equipotent to TAM on MDA-MB-468. Since TNBC cell lines do not express ER, this suggests that these novel TAM analogs elicit their anti-proliferative activity via a mechanism that does not involve binding to ER. The six

compounds also exhibited mild to high estrogenic activity, but with anti-proliferative activity, this offers an advantage over existing SERM such as TAM (Figure 2).

**Figure 2.** Dose-response curves of selected compounds on breast cancer cell lines. Data obtained from NCI in vitro disease-oriented human tumor cell screen (for details, see the work of [39]) compounds were tested in triplicates.

The ability of the compounds to inhibit the growth of other panels rather than breast cancer was investigated. All six compounds were found to be three times more active than TAM (mean GI<sup>50</sup> = 6.31 µM) on the colon cancer cell lines with (mean GI<sup>50</sup> = 1.90 µM). TAM was reported to inhibit the growth of colon cancer cells, yet the mechanism of inhibition is not clear yet, and further studies are warranted before any clinical implications can be postulated (see Supplementary Materials).

Compound **28** (mean GI<sup>50</sup> = 2.34 µM) was approximately three times as potent as TAM (mean GI<sup>50</sup> = 6.31 µM) on NSCLC cell lines, and twice as potent as TAM (mean GI<sup>50</sup> = 5.00 and 5.35 µM) on both renal (mean GI<sup>50</sup> = 2.40 µM) and prostate (mean GI<sup>50</sup> = 2.31 µM) cell lines. Compound **28** showed an exceptional broad-spectrum growth inhibition.

The six compounds showed the highest potency on colon cancer cell lines; this might indicate some selectivity toward this particular panel. Further investigations might help understand the reason for this selectivity (Figure 3).

**Figure 3.** Dose-response curves of compound **28** on different subpanels. Data obtained from NCI in vitro disease-oriented human tumor cell screen (for details, see the work of [39]) compounds were tested in triplicates.

α

#### *2.5. Alkaline Phosphatase Activity in Ishikawa Cell Line*

Because of the potential SERM character of the compounds tested, their estrogenic effects were studied in an endometrial-derived cell culture model, the human endometrial adenocarcinoma cell line Ishikawa. Estrogenic compounds are able to increase the alkaline phosphatase (AlkP) activity mediated by the ERα. All compounds were screened at two concentrations, 0.1 and 1 µM. Its agonistic effect was compared to the vehicle control DMSO (data shown in Supplementary Materials). Estradiol at 10 nM was used as a positive control and TAM and OH-TAM at 1 µM as comparative controls. Most of the compounds showed no significant increase in AlkP activity after a 72 h treatment. Compounds **5**, **11**, **12,** and **19** showed moderate estrogenic activity in YES assay and growth inhibition above 50% on MCF-7 cells at 10 µM; therefore, they were selected for the 5-dose AlkP assay. The four compounds were studied in a concentration range of 1 nM to 10 µM. Compounds **11**, **12**, and **19** were able to increase the AlkP activity in a dose-pendent manner with significant effects at a concentration of 100 nM and 1 µM. No significant effects were observed for compound **5**. The decreased activities at a concentration of 10 µM are caused by a negative influence of the treatment on the cell growth, observed with light microscopy. Compound **12** showed an equipotent activity when compared to TAM and 4-OH-TAM despite its higher relative estrogenic activity in the YES assay (Table 6).

**Table 6.** Relative alkaline phosphatase activity after an incubation of 72 h in Ishikawa cells.


Solvent control (DMSO) was set to 1 \* *p* < 0.05 (Tukey test) \*\* n.d. = not determined.

The observed moderate estrogenic effects of **11**, **12**, and **19** endorse the results obtained by the other in vitro assays reported. Using this Ishikawa cell culture model only gives a hint about possible effects on uterine tissue and needs more investigations.

#### *2.6. Uterotrophic Assay*

The most common short-term in vivo assay for estrogenicity/anti-estrogenicity is the uterotrophic assay, suitable for screening ERα agonists and antagonists. The primary endpoint is the uterine wet weight (UWW). An increase in UWW indicates an estrogenic activity of the test compound. Compounds **12** and **19** were screened using the in vivo uterotrophic assay. Both compounds showed less increase in UWW, indicating lower endometrial estrogenic activity and potentially less tendency to induce endometrial carcinoma (Table 7).

**Table 7.** Relative uterus wet weight of ovariectomized rats**.**


#### *2.7. In Silico Study*

The most potent estrogenic compound **3** (EC<sup>50</sup> = 40.1 nM) bearing an OH group at the para position of ring **B** and 3-fluoro 4-methoxy substituents on ring **A** was selected for the in

silico model. Compound **3** was docked into ERα LBD co-crystallized with diethylstilbestrol (DES), a synthetic estrogen with full agonistic activity (PDB: 3ERD) [40]. To validate the docking protocol, the co-crystallized ligand DES was docked into the ERα LBD where all the resultant poses converged to a similar binding mode as that of the experimentally determined position of DES with the best ranking pose having an RMSD value of 1.71 Å.

The crystal structures of ERα bound to DES (PDB code: 3ERD) [9] were downloaded from the PDB database. Only protein molecules were considered where it was optimized using the structure preparation wizard in MOE (version 2009.10) [38] and saved as a mol file. DES was built as E-isomer, whereas compound **3** was built as pure E and Z isomers, minimized using the MMFF94x force field in MOE using a gradient of 0.0001 kcal/(mol Å), and their protonation states at pH 7.0 were generated. A conformational search was adopted for compound **3E** and **3Z** isomers and E-DES. The database obtained was saved as.mdb and used as docking ligands.

Results of the overlay of compounds **3E** and **3Z** on DES showed that the **3E** conformer with the lowest binding energy showed a partial overlay on DES (Figure 4).

α **Figure 4.** Compound **3***E* (**cyan**) overlaid with DES (**yellow**) inside ERα LBD.

Compound **3E** retained the two essential interactions with Glu353 and His524, the oxygen of the methoxy group on ring **A** of compound **3E** acted as H-bond acceptor rather than H-bond donor (Figure 5).

α

α

α **Figure 5.** Two-dimensional interactions of DES (**red**) and compound **3E** (**green**) inside ERα LBD.

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

#### *3.1. Chemistry*

All reactions were carried out under nitrogen when an inert atmosphere was needed. Syntheses that required dry and oxygen-free conditions were performed in a Glovebox MB Unilab or using Schlenk techniques under an atmosphere of purified nitrogen or argon, respectively. Dry, oxygen-free solvents (CH2Cl2, distilled from CaH2; THF, distilled from potassium) were employed. All distilled and deuterated solvents were stored over molecular sieves (4 Å). All glassware was oven-dried at 160 ◦C prior to use. Solvents and reagents were obtained from commercial suppliers and were of pure analytical grade. Purification of compounds was carried out using column chromatography with silica gel 40– 60 µM mesh or using a Biotage*®* Isolera™ (Uppsala, Sweden) flash purification system using Biotage*®* KP-Sil SNAP columns. Reaction progress was monitored by TLC using fluorescent pre-coated silica gel plates, and detection of the components was made by short UV light (λ = 254 nm).

<sup>1</sup>H-NMR spectra were measured on either 400 MHz Bruker or on a Bruker AVANCE III HD Nanobay, 400 MHz UltraSield (1H (400.13 MHz), <sup>13</sup>C (100.61 MHz)) or on a Bruker AVANCE III HDX, 500 MHz Ascend (1H (500.13 MHz), <sup>13</sup>C (125.75 MHz)) spectrometer. All <sup>13</sup>C NMR spectra were exclusively recorded with composite pulse decoupling. Chemical shifts were referenced to δTMS = 0.00 ppm. (1H, <sup>13</sup>C) Chemical shifts (δ) are reported in ppm. Coupling constants (J) are reported in Hz. Multiplicities are abbreviated as: s: singlet; d: doublet; t: triplet; q: quartet; m: multiplet; dd: doublet of doublet; dt: doublet of triplet; brs: broad singlet. Mass spectrometric analysis (UPLC-ESI-MS) was performed using Waters ACQUITY Xevo TQD system, which consisted of an ACQUITY UPLC H-Class system and XevoTM TQD triple-quadrupole tandem mass spectrometer with an electrospray ionization (ESI) interface (Waters Corp., Milford, MA, USA). Acquity BEH C18 100 × 2.1 mm column (particle size, 1.7 µm) was used to separate analytes (Waters, Dublin, Ireland). The solvent system consisted of water containing 0.1% TFA (A) and 0.1% TFA in acetonitrile (B). UPLC-method: flow rate 200 µL/min. The percentage of B started at an initial of 5% and maintained for 1 min, then increased up to 100% during 10 min, kept at 100% for 2 min, and flushed back to 5% in 3 min. The MS scan was carried out at

the following conditions: capillary voltage 3.5 kV, cone voltage 20 V, radio frequency (RF) lens voltage 2.5 V, source temperature 150 ◦C, and desolvation gas temperature 500 ◦C. Nitrogen was used as the desolvation and cone gas at a flow rate of 1000 and 20 L/h, respectively. System operation and data acquisition were controlled using Mass Lynx 4.1 software (Waters).

#### 3.1.1. General Procedures for Preparation of Compound **1**–**4**

Zinc powder (10.11 g, 154 mmol) was suspended in dry THF (100 mL), and the mixture was cooled to 0 ◦C. TiCl<sup>4</sup> (7.5 mL, 70 mmol) was added dropwise under nitrogen/argon. When the addition was complete, the mixture was warmed to room temperature and heated to reflux for 2 h. After cooling down, a solution of 4-Chloro-4-hydroxybenzophenone (2.86 g, 12.3 mmol) and acetophenone/4′ -methoxyacetophenone/3′ -Fluoro-4′ -methoxyacetophen one/4′ -Fluoro-3′ -methoxyacetophenone (38.4 mmol) in dry THF (100 mL) was added at 0 ◦C, and the mixture was heated at reflux in the dark for 2.5–7 h. After being cooled to room temperature, the zinc dust was filtered off, and THF was removed under reduced pressure. The residue was dissolved in an aqueous solution containing 30% hydrochloric acid (500 mL) and then extracted with dichloromethane (120 mL × 6). The organic layers were combined and dried over anhydrous Na2SO4, concentrated in vacuo, and further purified by silica gel column chromatography or a Biotage*®* Isolera™ flash purification system using Biotage*®* KP-Sil SNAP columns (dichloromethane) to yield compounds **1**–**4** [34].

#### *E/Z*-4-[1-(4-Chloro-phenyl)-2-phenylpropenyl]-phenol (**1**)

C21H17ClO. Yield: 58%. Orange oil. Purity: 100%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ: 7.33 (d, J = 2.0 Hz, 2H), 7.32 (d, J = 2.0 Hz, 2H), 7.20–7.11 (m, 12H), 6.83 (d, J = 1.5 Hz, 1H), 6.80 (d, J = 1.5 Hz, 1H), 6.74 (d, J = 2.1 Hz, 2H), 6.72 (d, J = 2.1 Hz, 2H), 6.50 (d, J = 2.1 Hz, 2H), 6.49 (d, J = 2.1 Hz, 2H), 2.15 (s, 2H), 2.12 (s, 6H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 154.28, 153.60, 143.91, 143.82, 142.12, 141.78, 137.55, 137.51, 136.13, 135.67, 135.51, 135.34, 132.33, 132.18, 132.14, 131.52, 131.39, 129.19, 128.30, 128.03, 127.95, 127.58, 126.33, 126.23, 115.04, 114.45, 23.49, 23.30. MS (ESI): *m/z* = 321.1 [M+H]<sup>+</sup> (100%), *m/z* = 323.1 [M+H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.42 (100% methylene chloride).

#### *E/Z*-4-[1-(4-Chloro-phenyl)-2-(4-methoxy-phenyl)-propenyl]-phenol (**2**)

C22H19ClO2. Yield: 55%. Orange oil. Purity: 95%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ: 7.32 (d, J = 2.0 Hz, 1H), 7.30 (d, J = 1.9 Hz, 1H), 7.18 (d, J = 2.0 Hz, 1H), 7.16 (d, J = 1.8 Hz, 1H), 7.13–6.99 (m, 10H), 6.88 (d, J = 2.2 Hz, 1H), 6.87 (d, J = 2.1 Hz, 1H), 6.83 (d, J = 1.7 Hz, 2H), 6.81 (d, J = 1.8 Hz, 2H), 6.74–6.71 (m, 2H), 6.54 (d, J = 2.2 Hz, 1H), 6.52 (d, J = 2.0 Hz, 1H), 3.76 (d, J = 2.7 Hz, 6H), 2.58 (s, 2H), 2.12 (d, J = 5.0 Hz, 6H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 158.33, 157.94, 154.53, 153.80, 142.45, 142.15, 137.09, 137.02, 136.16, 136.07, 135.68, 135.65, 135.47, 135.39, 132.16, 132.12, 131.43, 131.31, 130.75, 130.36, 130.34, 130.15, 128.26, 127.61, 115.06, 114.55, 113.76, 113.72, 113.42, 113.34, 55.49, 55.15, 23.45, 23.24. MS (ESI): *m/z* = 351.1 [M+H]<sup>+</sup> (100%) *m/z* = 353.1 [M+H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.33 (100% methylene chloride).

#### *E/Z*-4-[1-(4-Chlorophenyl)-2-(3-fluoro-4-methoxyphenyl) propenyl]-phenol (**3**)

C22H18ClFO2. Yield: 70%. Orange oil. Purity: 95%. <sup>1</sup>H-NMR (500 MHz, CDCl3) (δ 7.33–7.29 (m, 2H), 7.16–7.13 (m, 2H), 7.08–7.05 (m, 2H), 7.04–7.00 (m, 2H), 6.90–6.88 (m, 1H), 6.88–6.85 (m, 1H), 6.83–6.81 (m, 3H), 6.81–6.79 (m, 2H), 6.77–6.76 (m, 1H), 6.76–6.73 (m, 2H), 6.73–6.71 (m, 2H), 6.55–6.51 (m, 2H), 5.30 (s, 2H), 3.84 (d, J = 1.7 Hz, 6H), 2.10 (s, 3H), 2.06 (s, 3H). <sup>13</sup>C-NMR (126 MHz, CDCl3) δ 154.47, 153.87, 151.89 (d, J = 245.2 Hz), 151.86 (d, J = 244.9 Hz), 145.94 (t, J = 10.7 Hz), 142.01, 141.70, 137.94, 136.89 (d, J = 6.3 Hz), 136.80 (d, J = 6.3 Hz), 135.45, 135.14, 134.43 (d, J = 1.4 Hz), 133.80 (d, J = 1.4 Hz), 132.42, 132.09, 132.04, 131.71, 131.33, 128.34, 127.78, 125.21 (d, J = 3.3 Hz), 125.13 (d, J = 3.3 Hz), 116.92 (d, J = 18.4 Hz), 116.86 (d, J = 18.3 Hz), 115.08, 114.66, 112.82 (d, J = 8.3 Hz), 112.81 (d, J = 8.3 Hz), 58.59, 56.16, 23.29, 23.09. MS (ESI): *m/z* = 368.83 [M+H]<sup>+</sup> , *m/z* = 370.83 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.45 (100% methylene chloride).

#### *E/Z*-4-[1-(4-Chlorophenyl)-2-(4-fluoro-3-methoxyphenyl) propenyl]-phenol (**4**)

C22H18ClFO2. Yield: 57%. Orange oil. Purity: 97%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.32 (d, J = 8.3 Hz, 2H), 7.17 (d, J = 8.3 Hz, 2H), 7.08 (d, J = 8.5 Hz, 2H), 7.03 (d, J = 8.4 Hz, 2H), 6.94–6.86 (m, 2H), 6.83 (t, J = 7.8 Hz, 3H), 6.78–6.68 (m, 5H), 6.65 (td, J = 8.2, 1.8 Hz, 2H), 6.54 (d, J = 8.5 Hz, 2H), 3.63 (d, J = 4.4 Hz, 6H), 2.19 (s, 4H), 2.12 (d, J = 13.8 Hz, 2H), 2.10 (s, 3H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ 154.49, 153.91, 151.00 (d, J = 245.6 Hz), 150.93 (d, J = 245.4 Hz), 146.88 (d, J = 10.8 Hz), 146.80 (d, J = 10.8 Hz), 141.88, 141.84, 140.01 (d, J = 4.0 Hz), 139.97 (d, J = 4.0 Hz), 138.01, 138.00, 135.31, 135.25, 135.09, 134.51, 132.47, 132.00, 131.95, 131.76, 131.31, 128.35, 127.77, 121.47 (d, J = 6.6 Hz), 121.36 (d, J = 6.6 Hz), 115.55 (d, J = 18.2 Hz), 115.44 (d, J = 18.3 Hz), 115.19, 115.16, 115.10, 114.64, 56.08, 56.04, 23.22, 23.00. MS (ESI): *m/z* = 368.83 [M+H]<sup>+</sup> (100%), *m/z* = 370.83 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.37 (100% methylene chloride).

#### 3.1.2. General Procedures for Preparation of Compounds **5**–**28**

A solution of compounds **1**–**4** (16.28 g, 47 mmol) in DMF (100 mL) was treated with K2CO<sup>3</sup> (19.5 g, 141 mmol) and heated in an oil bath at 80 ◦C. The resulting suspension was treated with the appropriate commercially available base hydrochloride salt (51 mmol) portion-wise over a 2 h period and stirred for 16 h. The reaction mixture was cooled to room temperature. K2CO<sup>3</sup> was filtered off, and DMF was removed under reduced pressure. The final product was purified by silica gel column chromatography or a Biotage*®* Isolera™ flash purification system using Biotage*®* KP-Sil SNAP columns (dichloromethane) to yield compounds (**5**–**28**).

#### *E/Z*-(3-{4-[1-(4-Chloro-phenyl)-2-phenyl-propenyl]-phenoxy}-propyl)-dimethyl-amine (**5**)

C26H28ClNO. Yield: 48%. Brown oil. Purity: 98%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ: 7.30 (d, J = 8.4 Hz, 2H), 7.19–7.09 (m, 16H), 6.97 (d, J = 8.5 Hz, 2H), 6.86 (d, J = 8.6 Hz, 4H), 6.80–6.73 (m, 2H), 4.06 (t, J = 5.9 Hz, 2H), 3.91 (t, J = 5.9 Hz, 2H), 2.90 (m, 2H), 2.83 (m, 2H), 2.59 (s, 6H), 2.55 (s, 6H), 2.18–2.16 (dd, J = 9.6, 6.1 Hz, 2H), 2.14–2.08 (m, 8H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 141.76, 135.78, 132.15, 131.97, 131.37, 131.23, 129.18, 129.16, 128.28, 128.00, 127.94, 127.55, 126.32, 126.22, 114.02, 113.38, 65.30, 56.11, 56.05, 44.18, 44.07, 25.95, 25.78, 23.48, 23.33. MS (ESI): *m/z* = 406.3 [M+H]<sup>+</sup> (100%), *m/z* = 408.2 [M+H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.43 (9:1 methylene chloride: methanol).

#### *E/Z*-1-(2-{4-[1-(4-Chloro-phenyl)-2-phenyl-propenyl]-phenoxy}-ethyl)-pyrrolidine (**6**)

C27H28ClNO. Yield: 44%. Faint brown oil. Purity: 95.84%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ: 7.33–7.29 (m, J = 2.0 Hz, 2H), 7.20–7.08 (m, 14H), 6.99–6.96 (m, 2H), 6.91–6.87 (m, 2H), 6.81–6.73 (m, 4H), 6.68–6.57 (m, 2H), 4.23 (t, J = 5.6 Hz, 2H), 4.09 (t, J = 5.6 Hz, 2H), 3.09 (t, J = 5.6 Hz, 2H), 3.01 (t, J = 5.5 Hz, 2H), 2.85 (d, J = 18.8 Hz, 8H), 2.13 (s, 3H), 2.10 (s, 3H), 1.96–1.84 (m, 8H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 157.17, 156.46, 143.84, 143.79, 142.13, 141.77, 137.52, 137.45, 136.14, 135.84, 135.51, 132.29, 132.18, 131.98, 131.48, 131.41, 131.23, 129.20, 129.17, 128.28, 128.01, 127.97, 127.56, 126.32, 126.25, 114.15, 113.51, 65.95, 65.65, 54.77, 54.69, 54.63, 54.58, 23.50, 23.39, 23.34. MS (ESI): *m/z* = 418.3 [M+H]<sup>+</sup> (100%), *m/z* = 420.3 [M+H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.5 (9:1 methylene chloride: methanol).

#### *E/Z*-1-(2-{4-[1-(4-Chloro-phenyl)-2-phenyl-propenyl]-phenoxy}-ethyl)-piperidine (**7**)

C28H30ClNO. Yield: 42%. Faint brown oil. Purity: 97.82%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ: 7.32 (d, J = 2.0 Hz, 1H), 7.30 (d, J = 1.6 Hz, 1H), 7.20–7.09 (m, 14H), 6.98 (d, J = 1.9 Hz, 1H), 6.97 (d, J = 2.0 Hz, 1H), 6.89 (d, J = 2.0 Hz, 1H), 6.87 (d, J = 2.0 Hz, 1H), 6.80 (d, J = 2.0 Hz, 1H), 6.79 (d, J = 1.9 Hz, 1H), 6.76 (d, J = 2.0 Hz, 1H), 6.74 (d, J = 2.1 Hz, 1H), 6.56 (d, J = 2.1 Hz, 1H), 6.55 (d, J = 2.0 Hz, 1H), 4.20 (t, J = 5.8 Hz, 2H), 4.05 (t, J = 5.7 Hz, 2H), 2.91 (t, J = 5.7 Hz, 2H), 2.82 (t, J = 5.7 Hz, 2H), 2.72–2.55 (m, 8H), 2.14 (s, 3H), 2.10 (s, 3H), 1.72–1.64

(m, 8H), 1.52–1.44 (m, 4H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 157.30, 156.61, 143.87, 143.80, 142.16, 141.79, 137.56, 137.49, 136.09, 135.69, 135.45, 135.34, 132.29, 132.18, 131.95, 131.48, 131.41, 131.21, 129.20, 129.18, 128.28, 128.01, 127.96, 127.56, 126.31, 126.23, 114.13, 113.52, 65.28, 65.03, 57.67, 57.58, 54.87, 54.80, 25.34, 25.23, 23.75, 23.68, 23.50, 23.33. MS (ESI): *m/z* = 432.3 [M+H]<sup>+</sup> (100%), *m/z* = 434.3 [M+H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.37 (93:7 methylene chloride: methanol).

#### *E/Z*-4-(2-{4-[1-(4-Chloro-phenyl)-2-phenyl-propenyl]-phenoxy}-ethyl)-morpholine (**8**)

C27H28ClNO2. Yield: 48%. Orange oil. Purity: 100%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ: 7.32 (d, J = 1.9 Hz, 1H), 7.31 (d, J = 2.0 Hz, 1H), 7.20–7.09 (m, 14H), 6.99 (d, J = 1.9 Hz, 1H), 6.97 (d, J = 2.0 Hz, 1H), 6.90 (d, J = 2.0 Hz, 1H), 6.88 (d, J = 2.0 Hz, 1H), 6.81 (d, J = 2.0 Hz, 1H), 6.79 (d, J = 1.9 Hz, 1H), 6.76 (d, J = 2.0 Hz, 1H), 6.75 (d, J = 2.1 Hz, 1H), 6.57 (d, J = 2.1 Hz, 1H), 6.56 (d, J = 2.0 Hz, 1H), 4.14 (t, J = 5.7 Hz, 2H), 4.00 (t, J = 5.7 Hz, 2H), 3.76 (m, 4H), 3.72 (m, 4H), 2.84 (t, J = 5.7 Hz, 2H), 2.74 (t, J = 5.7 Hz, 2H), 2.62 (m, 4H), 2.55 (m, 4H), 2.14 (s, 3H), 2.11 (s, 3H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 157.44, 156.78, 143.90, 143.80, 142.17, 141.80, 137.57, 137.51, 136.09, 135.67, 135.44, 135.28, 132.30, 132.18, 131.93, 131.49, 131.40, 131.20, 129.20, 129.19, 128.29, 128.02, 127.95, 127.56, 126.32, 126.21, 114.14, 113.53, 66.85, 66.80, 65.62, 65.41, 57.66, 57.62, 54.06, 54.02, 23.50, 23.34. MS (ESI): *m/z* = 434.3 [M+H]<sup>+</sup> (100%), *m/z* = 436.3 [M+H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.68 (95:5 methylene chloride: methanol).

#### *E/Z*-1-(2-{4-[1-(4-Chloro-phenyl)-2-phenyl-propenyl]-phenoxy}-ethyl)-azepane (**9**)

C29H32ClNO. Yield: 40%. Brown oil. Purity: 95.34%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ: 7.32 (d, J = 1.9 Hz, 1H), 7.30 (d, J = 1.9 Hz, 1H), 7.20–7.08 (m, 14H), 6.99–6.95 (m, 2H), 6.88 (dd, J = 6.7, 4.8 Hz, 2H), 6.80 (d, J = 1.9 Hz, 1H), 6.78 (d, J = 1.9 Hz, 1H), 6.76 (d, J = 2.0 Hz, 1H), 6.74 (d, J = 2.0 Hz, 1H), 6.56 (d, J = 1.9 Hz, 1H), 6.54 (d, J = 2.0 Hz, 1H), 4.21 (t, J = 5.7 Hz, 2H), 4.06 (t, J = 5.7 Hz, 2H), 3.12 (t, J = 5.7 Hz, 2H), 3.04 (t, J = 5.7 Hz, 2H), 3.00–2.89 (m, 8H), 2.14 (s, 3H), 2.10 (s, 3H), 1.80–1.58 (m, 16H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 156.76, 156.25, 143.78, 141.76, 136.13, 132.17, 131.97, 131.48, 131.40, 131.23, 129.19, 129.17, 128.29, 128.01, 127.96, 127.56, 126.31, 126.24, 114.15, 113.52, 56.32, 56.20, 55.69, 55.61, 26.96, 26.92, 26.57, 23.50, 23.33. MS (ESI): *m/z* = 446.3 [M+H]<sup>+</sup> (100%), *m/z* = 448.3 [M++H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.37 (93:7 methylene chloride: methanol).

#### *E/Z*-(3-{4-[1-(4-Chloro-phenyl)-2-(4-methoxy-phenyl)-propenyl]-phenoxy}-propyl) dimethyl-amine (**10**)

C27H30ClNO2. Yield: 48%. Orange oil. Purity: 96.57%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ: 7.30 (d, J = 1.9 Hz, 1H), 7.28 (d, J = 1.9 Hz, 1H), 7.16–6.96 (m, 12H), 6.87–6.68 (m, 8H), 6.56 (d, J = 2.1 Hz, 1H), 6.54 (d, J = 2.0 Hz, 1H), 4.05 (t, J = 6.0 Hz, 2H), 3.92 (t, J = 6.0 Hz, 2H), 3.75 (s, 6H), 2.84–2.73 (m, 4H), 2.51 (d, J = 10.2 Hz, 6H), 2.20–2.11 (m, 4H), 2.10 (s, 6H), 2.07 (s, 6H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 157.96, 157.87, 157.32, 156.59, 142.39, 142.08, 136.96, 136.88, 136.05, 135.96, 135.93, 135.65, 135.54, 134.89, 132.21, 131.99, 131.42, 131.29, 131.24, 130.33, 130.31, 128.26, 127.60, 113.99, 113.43, 113.37, 113.31, 65.42, 65.16, 56.21, 56.16, 55.13, 53.44, 44.47, 44.35, 26.28, 26.13, 23.43, 23.28. MS (ESI): *m/z* = 436.3 [M+H]<sup>+</sup> (100%), *m/z* = 438.3 [M+H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.45 (9:1 methylene chloride: methanol).

*E/Z*-1-(2-{4-[1-(4-Chloro-phenyl)-2-(4-methoxy-phenyl)-propenyl]-phenoxy} ethyl)-pyrrolidine (**10**)

C28H30ClNO2. Yield: 40%. Orange oil. Purity: 100%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ: 7.31 (d, J = 1.9 Hz, 1H), 7.29 (d, J = 1.9 Hz, 1H), 7.17–6.96 (m, 12H), 6.89 (d, J = 2.0 Hz, 1H), 6.87 (d, J = 2.0 Hz, 1H), 6.82–6.74 (m, 4H), 6.72 (d, J = 2.1 Hz, 1H), 6.70 (d, J = 2.1 Hz, 1H), 6.59 (d, J = 2.1 Hz, 1H), 6.57 (d, J = 2.0 Hz, 1H), 4.24 (t, J = 5.6 Hz, 2H), 4.12 (t, J = 5.6 Hz, 2H), 3.76 (s, 6H), 3.09 (t, J = 5.5 Hz, 2H), 3.03 (t, J = 5.4 Hz, 2H), 2.88 (s, 8H), 2.11 (s, 3H), 2.07 (s, 3H), 1.95–1.89 (m, 8H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 157.96, 157.89, 157.06, 142.06, 136.94, 136.12, 135.99, 135.94, 135.57, 132.22, 131.99, 131.43, 131.30, 131.25, 130.34, 130.31, 128.26, 127.61, 114.13, 113.56, 113.37, 113.33, 55.13, 54.78, 54.71, 54.64, 54.61, 23.44, 23.39, 23.35, 23.27. MS (ESI): *m/z* = 448.3 [M+H]<sup>+</sup> , *m/z* = 450.2 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.5 (9:1 methylene chloride: methanol).

*E/Z*-1-(2-{4-[1-(4-Chloro-phenyl)-2-(4-methoxy-phenyl)-propenyl]-phenoxy}-ethyl) piperidine (**12**)

C29H32ClNO2. Yield: 53%. Yellow oil. Purity: 100%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ: 7.31 (d, J = 2.0 Hz, 1H), 7.29 (d, J = 2.1 Hz, 1H), 7.18–6.98 (m, 10H), 6.88 (d, J = 2.2 Hz, 1H), 6.87 (d, J = 2.1 Hz, 1H), 6.83–6.69 (m, 8H), 6.59 (d, J = 2.2 Hz, 1H), 6.57 (d, J = 2.1 Hz, 1H), 4.15 (t, J = 6.0 Hz, 2H), 4.03 (t, J = 6.0 Hz, 2H), 3.76 (s, 6H), 2.84 (t, J = 6.0 Hz, 2H), 2.76 (t, J = 6.0 Hz, 2H), 2.56 (d, J = 18.9 Hz, 8H), 2.11 (s, 3H), 2.08 (s, 2H), 1.64 (tt, J = 11.6, 5.6 Hz, 9H), 1.47 (dd, J = 13.2, 8.4 Hz, 4H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 157.98, 157.89, 157.42, 156.73, 142.45, 142.12, 137.06, 136.97, 136.08, 136.00, 135.82, 135.49, 134.79, 132.21, 132.15, 131.91, 131.41, 131.30, 131.17, 130.33, 130.30, 128.25, 127.59, 114.15, 113.61, 113.38, 113.32, 65.65, 65.45, 57.86, 57.81, 55.12, 54.99, 54.95, 25.69, 25.62, 24.01, 23.95, 23.43, 23.25. MS (ESI): *m/z* = 462.3 [M+H]<sup>+</sup> (100%), *m/z* = 464.2 [M+H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.37 (93:7 methylene chloride: methanol).

*E/Z*-4-(2-{4-[1-(4-Chloro-phenyl)-2-(4-methoxy-phenyl)-propenyl]-phenoxy}-ethyl) morpholine (**13**)

C28H30ClNO3. Yield: 45%. Dark orange oil. Purity: 97.45%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.31 (d, J = 1.7 Hz, 1H), 7.29 (d, J = 2.0 Hz, 1H), 7.17–6.97 (m, 10H), 6.89 (d, J = 2.8 Hz, 1H), 6.87 (d, J = 2.0 Hz, 1H), 6.82–6.74 (m, 4H), 6.73–6.68 (m, 4H), 6.59 (d, J = 1.9 Hz, 1H), 6.57 (d, J = 2.0 Hz, 1H), 4.14 (t, J = 5.7 Hz, 2H), 4.01 (t, J = 5.7 Hz, 2H), 3.83–3.69 (m, 14H), 2.83 (t, J = 5.6 Hz, 2H), 2.75 (t, J = 5.6 Hz, 2H), 2.59 (d, J = 20.5 Hz, 8H), 2.11 (s, 3H), 2.08 (s, 3H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ: 157.97, 157.87, 157.36, 156.68, 142.42, 142.10, 136.99, 136.92, 136.07, 135.94, 135.92, 135.59, 135.52, 134.85, 132.22, 132.16, 131.95, 131.43, 131.31, 131.21, 130.34, 130.32, 128.26, 127.61, 114.12, 113.58, 113.37, 113.31, 66.86, 65.61, 65.42, 57.67, 57.65, 55.13, 54.06, 54.04, 53.43, 23.44, 23.28. MS (ESI): *m/z* = 464.3 [M+H]<sup>+</sup> (100%), *m/z* = 466.2 [M+H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.52 (95:5 methylene chloride: methanol).

*E/Z*-1-(2-{4-[1-(4-Chloro-phenyl)-2-(4-methoxy-phenyl)-propenyl]-phenoxy}-ethyl) azepane (**14**)

C30H34ClNO2. Yield: 43%. Orange oil. Purity: 95%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.32–7.25 (m, 2H), 7.17–7.08 (m, 4H), 7.06–6.97 (m, 6H), 6.87 (d, J = 8.7 Hz, 2H), 6.81–6.74 (m, 4H), 6.70 (d, J = 8.7 Hz, 4H), 6.59–6.55 (m, 2H), 4.19 (t, J = 5.6 Hz, 2H), 4.07 (t, J = 5.7 Hz, 2H), 3.76 (s, 6H), 3.09 (t, J = 5.6 Hz, 2H), 3.04–3.00 (m, 2H), 2.97–2.84 (m, 8H), 2.10 (s, 3H), 2.07 (s, 3H), 1.81–1.69 (m, 8H), 1.68–1.60 (m, 8H). <sup>13</sup>C-NMR (101 MHz, CDCl3) δ 157.94, 142.40, 142.07, 135.56, 134.91, 132.19, 131.96, 131.51–131.06, 130.31, 128.26, 127.60, 114.16, 113.60, 113.36, 56.25, 55.62, 55.12, 26.97, 23.34. MS (ESI): *m/z* = 476.4 [M+H]<sup>+</sup> (100%), *m/z* = 478.4 [M+H+2]<sup>+</sup> (33%). R<sup>f</sup> : 0.53 (9:1 methylene chloride: methanol).

*E/Z*-(3-{4-[1-(4-Chlorophenyl)-2-(3-fluoro-4-methoxyphenyl)-propenyl]-phenoxy} propyl)-dimethyl-amine (**15**)

C27H29ClFNO2. Yield: 74%. Orange oil. Purity: 98.84%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.31–7.28 (m, 2H), 7.15–7.12 (m, 2H), 7.10–7.07 (m, 2H), 7.02–6.99 (m, 2H), 6.88 (s, 2H), 6.85 (d, J = 2.9 Hz, 2H), 6.82–6.79 (m, 3H), 6.77–6.73 (m, 5H), 6.58 (dd, J = 9.1, 2.3 Hz, 2H), 4.02 (t, J = 6.4 Hz, 2H), 3.91 (t, J = 6.4 Hz, 2H), 3.83 (s, 6H), 3.46–3.33 (m, 4H), 2.25 (dd, J = 12.1, 5.9 Hz, 10H), 2.10–2.04 (m, 12H). <sup>13</sup>C-NMR: (101 MHz, CDCl3) δ 157.85, 157.27, 145.95 (d, J = 10.5 Hz), 145.92 (d, J = 10.5 Hz), 142.10, 141.75, 138.02, 137.95, 137.99, (d, J = 7.1 Hz), 135.18, 134.82, 134.29 (d, J = 1.4 Hz), 133.60 (d, J = 1.3 Hz), 132.32, 132.07, 131.78, 131.62, 131.30, 128.26, 127.71, 125.18 (d, J = 3.1 Hz), 125.08 (d, J = 3.1 Hz), 116.88 (d, J = 18.1 Hz), 116.83 (d, J = 18.3 Hz), 114.11, 113.66, 112.80 (d, J = 6.0 Hz), 112.78 (d, J = 5.9 Hz), 66.11, 65.93, 56.35 (d, J = 3.8 Hz), 56.10, 45.40, 45.35, 27.49, 27.41, 23.25, 23.06. MS (ESI): *m/z* = 454.30 [M+H]<sup>+</sup> , *m/z* = 456.30 [M+H+2]<sup>+</sup> (100%). R<sup>f</sup> : 0.41 (93:7 methylene chloride: methanol).

*E/Z*-1-(2-{4-[1-(4-Chlorophenyl)-2-(3-fluoro-4-methoxyphenyl)-propenyl]-phenoxy} ethyl)-pyrrolidine (**16**)

C28H29ClFNO2. Yield: 61%. Orange oil. Purity: 100%. <sup>1</sup>H-NMR (500 MHz, CDCl3) δ 7.31–7.27 (m, 2H), 7.15–7.10 (m, 2H), 7.10–7.06 (m, 2H), 7.02–6.98 (m, 2H), 6.88 (d, J = 2.1 Hz, 1H), 6.86 (t, J = 2.6 Hz, 2H), 6.84 (dd, J = 5.9, 3.5 Hz, 1H), 6.81–6.79 (m, 2H), 6.78 (t, J = 4.8 Hz, 2H), 6.76–6.71 (m, 4H), 6.62–6.57 (m, 2H), 4.15 (t, J = 5.9 Hz, 2H), 4.04 (t, J = 5.9 Hz, 2H), 3.82 (s, 6H), 2.65 (d, J = 20.4 Hz, 8H), 2.67–2.64 (m, 8H), 2.10 (s, 3H), 2.07 (s, 3H), 1.90–1.76 (m, 8H). <sup>13</sup>C-NMR: (126 MHz, CDCl3) δ 157.64, 157.06, 151.86 (d, J = 245.2 Hz), 151.84 (d, J = 244.9 Hz), 145.93 (t, J = 11.1 Hz), 142.08, 141.74, 138.00, 137.93, 136.87 (d, J = 6.4 Hz), 136.80 (d, J = 6.3 Hz), 135.38, 135.03, 134.36 (d, J = 1.3 Hz), 133.70 (d, J = 1.3 Hz), 132.36, 132.09, 131.80, 131.66, 131.32, 131.09, 128.30, 127.74, 125.20 (d, J = 3.3 Hz), 125.14 (d, J = 3.3 Hz), 116.87 (d, J = 18.4 Hz), 116.85 (d, J = 18.4 Hz), 114.19, 113.76, 112.80 (d, J = 5.9 Hz), 112.79 (d, J = 5.9 Hz), 66.82, 66.63, 56.12, 55.02 (d, J = 5.1 Hz), 54.70, 54.68, 23.48, 23.45, 23.28, 23.08. MS (ESI): *m/z* = 466.40 [M+H]<sup>+</sup> (100%), *m/z* = 468.00 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.34 (95:5 methylene chloride: methanol).

*E/Z*-1-(2-{4-[1-(4-Chlorophenyl)-2-(3-fluoro-4-methoxyphenyl)-propenyl]-phenoxy} ethyl)-piperidine (**17**)

C29H31ClFNO2. Yield: 77%. Orange oil. Purity: 95.88%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.30–7.27 (m, 2H), 7.14–7.11 (m, 2H), 7.08 (d, J = 8.7 Hz, 2H), 7.02–6.98 (m, 2H), 6.88–6.84 (m, 4H), 6.82–6.77 (m, 4H), 6.76–6.73 (m, 4H), 6.60–6.56 (m, 2H), 4.12 (t, J = 6.0 Hz, 2H), 4.00 (t, J = 6.0 Hz, 2H), 3.82 (s, 6H), 2.79 (t, J = 6.0 Hz, 2H), 2.72 (t, J = 6.0 Hz, 2H), 2.58 (m, 8H), 2.07 (s, 3H), 2.05 (s, 3H), 1.60 (dd, J = 11.3, 5.6 Hz, 8H), 1.46–1.40 (m, 4H). <sup>13</sup>C-NMR: (101 MHz, CDCl3) δ 157.65, 157.08, 151J = 10.8 Hz.87 (d, J = 245.2 Hz), 151.84 (d, J = 244.9 Hz), 145.97 (d, ), 145.88 (d, J = 10.8 Hz), 142.08, 141.74, 138.01, 137.94, 136.88 (d, J = 8.5 Hz), 136.82 (d, J = 8.4 Hz), 135.34, 134.98, 134.35, 133.68, 132.36, 132.08, 131.79, 131.66, 131.31, 131.07, 128.30, 127.74, 125.19 (d, J = 3.4 Hz), 125.11 (d, J = 3.3 Hz), 116.88 (d, J = 18.4 Hz), 116.85 (d, J = 18.3 Hz) 114.20, 113.77, 112.82 (d, J = 5.0 Hz), 112.80 (d, J = 5.1 Hz), 65.72 (d, J = 17.0 Hz), 57.89 (d, J = 4.5 Hz), 56.12, 55.02, 54.99, 25.82, 25.76, 24.11, 24.07, 23.27, 23.08. MS (ESI): *m/z* = 480.01 [M+H]<sup>+</sup> , *m/z* = 482.01 [M+H+2]<sup>+</sup> (100%). R<sup>f</sup> : 0.30 (94:6 methylene chloride: methanol).

*E/Z*-4-(2-{4-[1-(4-Chlorophenyl)-2-(3-fluoro-4-methoxyphenyl)-propenyl]-phenoxy} ethyl)-morpholine (**18**)

C28H29ClFNO3. Yield: 65%. Brown oil. Purity: 100%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.32–7.25 (m, 3H), 7.16–7.07 (m, 4H), 7.03–6.99 (m, 2H), 6.90–6.73 (m, 11H), 6.62–6.57 (m, 2H), 4.11 (t, J = 5.6 Hz, 2H), 3.99–4.02 (t, J = 5.7 Hz, 2H), 3.84 (s, 6H), 3.77–3.70 (m, 8H), 2.80 (t, J = 5.6 Hz, 2H), 2.73 (t, J = 5.6 Hz, 2H), 2.64–2.50 (m, 8H), 2.09 (s, 3H), 2.06 (s, 3H).13C-NMR: (101 MHz, CDCl3) δ 157.51, 156.93, 151.82 (d, J = 245.2 Hz), 151.79 (d, J = 244.8 Hz), 145.87 (d, J = 10.3 Hz), 142.02, 141.69, 137.91, 137.85, 136.83 (d, J = 6.2 Hz), 136.71 (d, J = 6.2 Hz), 135.47, 135.11, 134.40 (d, J = 1.3 Hz), 133.76 (d, J = 1.3 Hz), 132.34, 132.04, 131.78, 131.63, 131.28, 131.07, 128.27, 127.71, 125.15 (d, J = 3.3 Hz), 125.04 (d, J = 3.4 Hz), 116.88 (d, J = 18.4 Hz), 116.80 (d, J = 18.5 Hz), 114.18, 113.75, 112.80 (d, J = 5.4 Hz), 112.78 (d, J = 5.5 Hz), 66.80, 66.76, 65.65, 65.48, 57.62, 57.58, 57.60 (d, J = 3.3 Hz), 54.03, 54.00, 53.51, 23.22, 23.04. MS (ESI): *m/z* = 482.00 [M+H]<sup>+</sup> (100%), *m/z* = 484.00 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.58 (95:5 methylene chloride: methanol).

*E/Z*-1-(2-{4-[1-(4-Chlorophenyl)-2-(3-fluoro-4-methoxyphenyl)-propenyl]-phenoxy} ethyl)-azepane (**19**)

C30H33ClFNO2. Yield: 54%. Brown oil. Purity: 100%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.30–7.27 (m, 2H), 7.14–7.05 (m, 4H), 7.02–6.98 (m, 2H), 6.88–6.82 (m, 4H), 6.81–6.77 (m, 4H), 6.76–6.71 (m, 4H), 6.61 (dd, J = 6.8, 4.8 Hz, 2H), 4.18 (t, J = 6.0 Hz, 2H), 4.04 (t, J = 6.0 Hz, 2H), 3.83 (s, 6H), 3.03 (t, J = 6.0 Hz, 2H), 2.97 (t, J = 6.0 Hz, 2H), 2.90–2.81 (m, 8H), 2.08 (d, J = 14.2 Hz, 8H), 1.72 (d, J = 4.7 Hz, 6H), 1.65–1.60 (m, 8H). <sup>13</sup>C-NMR: (101 MHz, CDCl3) δ 158.79 (d, J =157.7 Hz), 157.56, 151.02, 150.83, 145.97, 144.90, 141.71, 136.39 (d, J =

6.2 Hz), 136.26 (d, J = 6.2 Hz), 135.71, 135.43, 134.38 (d, J = 1.3 Hz), 134.27 (d, J = 1.3 Hz), 132.37, 132.07, 131.81, 131.66, 131.30, 131.09, 128.29, 127.73, 125.18 (d, J = 3.1 Hz), 125.09 (d, J = 3.1 Hz), 116.93 (d, J = 18.4 Hz), 116.74 (d, J = 18.4 Hz), 114.21, 113.76, 112.82 (d, J = 5.7 Hz), 112.79 (d, J = 5.7 Hz), 66.06, 56.37, 56.11, 55.77 (d, J = 5.87 Hz), 53.41, 27.11, 27.01, 26.99, 26.90, 23.26, 23.07. MS (ESI): *m/z* = 494.04 [M+H]<sup>+</sup> (100%), *m/z* = 496.04 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.34 (93:7 methylene chloride: methanol).

*E/Z*-(2-{4-[1-(4-Chlorophenyl)-2-(3-fluoro-4-methoxyphenyl)-propenyl]-phenoxy} ethyl)-dimethyl-amine (**20**)

C26H27ClFNO2. Yield: 55%. Reddish-brown oil. Purity: 98.69%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.32–7.28 (m, 2H), 7.16–7.12 (m, 2H), 7.11–7.07 (m, 2H), 7.03–6.99 (m, 2H), 6.91–6.87 (m, 3H), 6.86 (d, J = 1.9 Hz, 1H), 6.82 (d, J = 1.9 Hz, 1H), 6.79 (dd, J = 5.7, 2.9 Hz, 2H), 6.78–6.74 (m, 4H), 6.72 (dd, J = 8.5, 1.5 Hz, 1H), 6.64–6.59 (m, 2H), 4.10 (t, J = 5.7 Hz, 2H), 3.98 (t, J = 5.7 Hz, 2H), 3.84 (s, 6H), 2.77 (t, J = 5.7 Hz, 2H), 2.70 (t, J = 5.7 Hz, 2H), 2.37 (s, 6H), 2.33 (s, 6H), 2.09 (s, 3H), 2.06 (s, 3H). <sup>13</sup>C-NMR: (101 MHz, CDCl3) δ 157.65, 157.07, 151.97 (d, J = 245.1 Hz), 151.94 (d, J = 244.9 Hz), 145.99 (d, J = 9.6 Hz), 142.07, 141.73, 138.01, 137.95, 136.81 (d, J = 6.7 Hz), 136.77 (d, J = 6.7 Hz),135.42, 135.07, 134.37 (d, J = 1.3 Hz), 133.72 (d, J = 1.6 Hz), 132.38, 132.09, 131.79, 131.68, 131.32, 131.09, 128.31, 127.76, 125.20 (d, J = 3.4 Hz), 125.13 (d, J = 3.4 Hz), 116.86 (d, J = 18.4 Hz), 116.98 (d, J = 18.3 Hz), 114.19, 113.75, 112.82 (d, J = 5.1 Hz), 112.80 (d, J = 5.2 Hz), 65.85, 65.64, 58.27, 58.22, 56.14, 45.83, 45.80, 23.29, 23.07. MS (ESI): *m/z* = 440.30 [M+H]<sup>+</sup> (100%), *m/z* = 442.30 [M+2]<sup>+</sup> . R<sup>f</sup> : 0.38 (92:8 methylene chloride: methanol).

*E/Z*-(2-{4-[1-(4-Chlorophenyl)-2-(3-fluoro-4-methoxyphenyl)-propenyl]-phenoxy} ethyl)-diethyl-amine (**21**)

C28H31ClFNO2. Yield: 51%. Yellow oil. Purity: 98.96%. <sup>1</sup>H-NMR (500 MHz, CDCl3) δ 7.30 (d, J = 8.4 Hz, 2H), 7.14 (d, J = 8.4 Hz, 2H), 7.09 (d, J = 8.6 Hz, 2H), 7.02 (d, J = 8.5 Hz, 2H), 6.87 (dd, J = 11.8, 2.6 Hz, 4H), 6.80 (t, J = 8.0 Hz, 3H), 6.78–6.70 (m, 5H), 6.59 (d, J = 8.7 Hz, 2H), 4.10 (d, J = 5.4 Hz, 2H), 4.00 (s, 2H), 3.84 (s, 6H), 2.95 (d, J = 4.8 Hz, 2H), 2.88 (s, 2H), 2.78–2.57 (m, 8H), 2.09 (d, J = 7.5 Hz, 3H), 2.09 (d, J = 7.5 Hz, 3H), 1.14–1.04 (m, 12H). <sup>13</sup>C-NMR: (126 MHz, CDCl3) δ 157.58, 156.96, 151.89 (d, J = 245.2 Hz), 151.85 (d, J = 244.8 Hz), 145.94 (t, J = 11.3 Hz), 142.07, 141.73, 137.99, 137.93, 136.89 (d, J = 6.2 Hz), 136.81 (d, J = 6.4 Hz), 135.43, 135.09, 134.38 (d, J = 1.1 Hz), 133.74 (d, J = 1.1 Hz), 132.39, 132.10, 131.84, 131.69, 131.33, 131.12, 128.32, 127.77, 125.21 (d, J = 3.3 Hz), 125.13 (d, J = 3.4 Hz), 116.91 (d, J = 18.4 Hz), 116.86 (d, J = 18.4 Hz), 114.15, 113.71, 112.81 (d, J = 5.9 Hz), 112.79 (d, J = 5.9 Hz), 66.12, 56.14, 53.42, 51.67, 51.54, 47.80, 47.77, 23.30, 23.09, 11.56. MS (ESI): *m/z* = 468.30 [M+H]<sup>+</sup> (100%), *m/z* = 470.00 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.32 (93:7 methylene chloride: methanol).

*E/Z*-(3-{4-[1-(4-Chlorophenyl)-2-(4-fluoro-3-methoxyphenyl)-propenyl]-phenoxy} propyl)-dimethyl-amine (**22**)

C27H29ClFNO2. Yield: 50%. Orange oil. Purity: 97.83%. <sup>1</sup>H-NMR (500 MHz, CDCl3) δ 7.31 (d, J = 8.4 Hz, 1H), 7.16 (d, J = 8.3 Hz, 1H), 7.10 (t, J = 5.7 Hz, 3H), 7.02 (d, J = 8.4 Hz, 3H), 6.92–6.85 (m, 5H), 6.81 (t, J = 7.1 Hz, 3H), 6.75 (d, J = 8.7 Hz, 1H), 6.72–6.67 (m, 2H), 6.65 (dd, J = 8.4, 1.9 Hz, 1H), 6.63 (dd, J = 8.3, 1.9 Hz, 1H), 6.59 (d, J = 8.7 Hz, 1H), 4.04 (t, J = 6.3 Hz, 2H), 3.91 (t, J = 6.3 Hz, 2H), 3.62 (d, J = 2.0 Hz, 6H), 2.56 (dt, J = 27.7, 7.4 Hz, 4H), 2.37–2.31 (m, 12H), 2.11 (d, J = 17.4 Hz, 6H), 2.07–1.94 (m, 4H). <sup>13</sup>C-NMR: (126 MHz, CDCl3) δ 157.79, 157.21, 150.97 (d, J = 245.6 Hz), 150.89 (d, J = 245.3 Hz), 146.89 (d, J = 10.8 Hz), 146.82 (d, J = 10.7 Hz), 141.93, 141.88, 140.08 (d, J = 4.0 Hz), 139.98 (d, J = 4.0 Hz), 138.09, 138.07, 135.19, 135.11, 135.02, 134.44, 132.44, 132.01, 131.75, 131.73, 131.33, 131.12, 128.34, 127.75, 121.42 (d, J = 6.6 Hz), 115.54 (d, J = 18.2 Hz), 115.43 (d, J = 18.2 Hz), 115.09, 114.13, 113.67, 65.94, 65.80, 56.37, 56.31, 56.03 (d, J = 2.6 Hz), 53.43, 45.17, 45.10, 27.16, 27.04, 23.24, 23.05.MS (ESI): *m/z* = 454.30 [M+H]<sup>+</sup> (100%), *m/z* = 456.30 [M++H+2]<sup>+</sup> . R<sup>f</sup> : 0.33 (91:9 methylene chloride: methanol).

*E/Z*-1-(2-{4-[1-(4-Chlorophenyl)-2-(4-fluoro-3-methoxyphenyl)-propenyl]-phenoxy} ethyl)-pyrrolidine (**23**)

C28H29ClFNO2. Yield: 71%. Brown oil. Purity: 100%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.31 (d, J = 8.4 Hz, 2H), 7.16 (d, J = 8.4 Hz, 2H), 7.11 (d, J = 8.6 Hz, 2H), 7.02 (d, J = 8.4 Hz, 2H), 6.89 (dt, J = 8.2, 5.5 Hz, 4H), 6.81 (d, J = 8.4 Hz, 2H), 6.76 (d, J = 8.7 Hz, 2H), 6.72–6.67 (m, 3H), 6.64 (t, J = 2.0 Hz, 1H), 6.61 (d, J = 8.6 Hz, 2H), 4.18 (t, J = 5.8 Hz, 2H), 4.05 (t, J = 5.8 Hz, 2H), 3.62 (d, J = 2.8 Hz, 6H), 2.98 (t, J = 5.8 Hz, 2H), 2.90 (t, J = 5.8 Hz, 2H), 2.71 (d, J = 21.8 Hz, 8H), 2.13 (s, 3H), 2.09 (s, 3H), 1.89–1.80 (m, 8H). <sup>13</sup>C-NMR: (101 MHz, CDCl3) δ 157.58, 157.01, 152.21, 152.16, 146.90 (d, J = 9.9 Hz), 146.89 (d, J = 9.8 Hz), 141.91, 141.86, 140.03 (d, J = 3.9 Hz), 139.97 (d, J = 4.1 Hz), 138.06, 138.03, 135.34, 135.25, 135.06, 134.47, 132.44, 132.00, 131.73, 131.31, 131.11, 128.33, 127.75, 121.45 (d, J = 6.0 Hz), 115.57 (d, J = 18.2 Hz), 115.39, (d, J = 18.3 Hz), 115.12 (d, J = 2.0 Hz), 115.11, 114.23, 113.79, 66.63, 66.50, 56.03 (d, J = 1.8 Hz), 54.99, 54.91, 54.69, 54.65, 23.48, 23.44, 23.22, 23.03. MS (ESI): *m/z* = 466.30 [M+H]<sup>+</sup> (100%), *m/z* = 468.20 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.32 (94:6 methylene chloride: methanol).

*E/Z*-1-(2-{4-[1-(4-Chlorophenyl)-2-(4-fluoro-3-methoxyphenyl)-propenyl]-phenoxy} ethyl)-piperidine (**24**)

C29H31ClFNO2. Yield: 42%. Orange oil. Purity: 95%. <sup>1</sup>H-NMR (400 MHz, CDCl3)) δ 7.33 (d, J = 7.5 Hz, 2H), 7.16 (dd, J = 17.6, 7.6 Hz, 4H), 7.04 (d, J = 7.6 Hz, 2H), 6.91 (t, J = 9.5 Hz, 4H), 6.83 (d, J = 7.4 Hz, 2H), 6.77 (d, J = 7.7 Hz, 2H), 6.70 (d, J = 13.9 Hz, 3H), 6.62 (d, J = 8.6 Hz, 3H), 4.23 (s, 2H), 4.10 (s, 2H), 3.66 (s, 6H), 2.91 (dd, J = 30.7, 19.6 Hz, 4H), 2.63 (s, 6H), 2.13 (d, J = 13.6 Hz, 6H), 1.71 (s, 8H), 1.47 (d, J = 23.0 Hz, 6H). <sup>13</sup>C-NMR: (101 MHz, CDCl3) δ 157.79, 157.21, 150.97, 150.89, 146.90 (d, J = 9.2 Hz), 146.81 (d, J = 9.0 Hz), 141.93, 141.88, 140.08 (d, J = 4.0 Hz), 139.98 (d, J = 4.0 Hz), 138.09, 138.07, 135.19, 135.11, 135.02, 134.44, 132.44, 132.01, 131.75, 131.73, 131.33, 131.12, 128.34, 127.75, 121.42 (t, J = 6.6 Hz), 115.56 (d, J = 13.4 Hz), 115.41 (d, J = 13.3 Hz), 115.09,114.33, 114.13, 113.67, 66.22, 57.98, 57.67, 56.04, 54.85 (d, J = 6.4 Hz), 53.40, 26.91, 25.63, 25.29, 23.72, 23.22, 23.04. MS (ESI): *m/z* = 480.01 [M+H]<sup>+</sup> (100%), *m/z* = 482.01 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.40 (95:5 methylene chloride: methanol).

*E/Z*-4-(2-{4-[1-(4-Chlorophenyl)-2-(4-fluoro-3-methoxyphenyl)-propenyl]-phenoxy} ethyl)-morpholine (**25**)

C28H29ClFNO3. Yield: 61%. Orange oil. Purity: 97.07%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.31 (d, J = 8.4 Hz, 2H), 7.14 (dd, J = 15.7, 8.5 Hz, 4H), 7.02 (d, J = 8.4 Hz, 2H), 6.90 (t, J = 9.1 Hz, 4H), 6.81 (d, J = 8.4 Hz, 2H), 6.76 (d, J = 8.7 Hz, 2H), 6.73–6.65 (m, 3H), 6.64 (dd, = 5.4, 3.3 Hz, 1H), 6.61 (t, J = 6.2 Hz, 2H), 4.15 (t, J = 5.5 Hz, 2H), 4.03 (t, J = 5.5 Hz, 2H), 3.80–3.70 (m, 8H), 3.63 (s, 6H), 2.86 (t, J = 5.3 Hz, 2H), 2.77 (t, J = 5.3 Hz, 2H), 2.61 (d, J = 21.5 Hz, 8H), 2.13 (s, 3H), 2.10 (s, 3H). <sup>13</sup>C-NMR: (101 MHz, CDCl3) δ 157.67, 157.09, 151.14 (d, J = 245.8 Hz), 151.06 (d, J = 245.2 Hz), 147.06 (d, J = 10.8 Hz), 146.99 (d, J = 10.8 Hz), 142.02, 141.98, 140.19 (d, J = 4.1 Hz), 140.06 (d, J = 4.0 Hz), 138.16, 138.14, 135.58, 135.49, 135.25, 134.70, 132.61, 132.13, 131.90, 131.44, 131.28, 128.49, 127.91, 121.57 (d, J = 5.1 Hz), 121.51 (d, J = 5.2 Hz), 115.69 (d, J = 18.2 Hz), 115.59 (d, J = 18.3 Hz), 115.27 (d, J = 1.4 Hz), 115.22 (d, J = 1.8 Hz), 115.12, 114.37, 113.93, 66.89, 65.70, 65.63, 57.76, 56.19, 54.19, 54.15, 53.56, 23.36, 23.19. MS (ESI): *m/z* = 482.30 [M+H]<sup>+</sup> (100%), *m/z* = 484.20 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.35 (94:6 methylene chloride: methanol).

*E/Z*-1-(2-{4-[1-(4-Chlorophenyl)-2-(4-fluoro-3-methoxyphenyl)-propenyl]-phenoxy} ethyl)-azepane (**26**)

C30H33ClFNO2. Yield: 62%. Orange oil. Purity: 95%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.31 (d, J = 8.4 Hz, 2H), 7.16 (d, J = 8.4 Hz, 2H), 7.11 (d, J = 8.6 Hz, 2H), 7.02 (d, J = 8.5 Hz, 2H), 6.93–6.86 (m, 4H), 6.81 (d, J = 8.4 Hz, 2H), 6.76 (d, J = 8.7 Hz, 2H), 6.73–6.66 (m, 3H), 6.64 (s, 1H), 6.61 (t, J = 6.3 Hz, 2H), 4.13 (t, J = 6.0 Hz, 2H), 4.00 (t, J = 6.0 Hz, 2H), 3.62 (d, J = 3.2 Hz, 6H), 3.01 (t, J = 6.0 Hz, 2H), 2.93 (t, J = 6.0 Hz, 2H), 2.87–2.76 (m, 8H), 2.13 (s, 3H), 2.10 (s, 3H), 1.74–1.65 (m, 8H), 1.64–1.58 (m, 8H). <sup>13</sup>C-NMR: (101 MHz, CDCl3)

δ 157.68, 157.10, 150.98 (d, J = 245.8 Hz), 150.91 (d, J = 245.4 Hz), 146.90 (d, J = 10.6 Hz), 146.83 (d, J = 10.8 Hz), 141.93, 141.87, 140.05 (d, J = 4.0 Hz), 139.97 (d, J = 4.0 Hz), 138.08, 138.06, 135.26, 135.17, 135.04, 134.45, 132.44, 132.00, 131.73, 131.31, 131.10, 128.33, 127.75, 121.42 (d, J = 6.9 Hz), 121.31, 115.53 (d, J = 18.4 Hz), 115.43 (d, J = 18.3 Hz), 115.12, 114.24, 113.80,66.16, 66.00, 56.40, 56.30, 56.03 (d, J = 2.3 Hz), 55.83, 55.77, 27.51, 27.47, 27.04, 27.02, 23.23, 23.03. MS (ESI): *m/z* = 494.04 [M+H]<sup>+</sup> (100%), *m/z* = 496.04 [M+2]<sup>+</sup> . R<sup>f</sup> : 0.50 (95:5 methylene chloride: methanol).

*E/Z*-(2-{4-[1-(4-Chlorophenyl)-2-(4-fluoro-3-methoxyphenyl)-propenyl]-phenoxy} ethyl)-dimethyl-amine (**27**)

C26H27ClFNO2. Yield: 67%. Yellow oil. Purity: 100%. <sup>1</sup>H-NMR (400 MHz, CDCl3) δ 7.31 (d, J = 8.4 Hz, 2H), 7.16 (d, J = 8.4 Hz, 2H), 7.11 (d, J = 8.6 Hz, 2H), 7.02 (d, J = 8.5 Hz, 2H), 6.92–6.86 (m, 4H), 6.81 (d, J = 8.5 Hz, 2H), 6.76 (d, J = 8.7 Hz, 2H), 6.73–6.65 (m, 3H), 6.62 (t, J = 7.5 Hz, 3H), 4.11 (t, J = 5.7 Hz, 2H), 3.98 (t, J = 5.7 Hz, 2H), 3.62 (d, J = 4.2 Hz, 6H), 2.78 (t, J = 5.7 Hz, 2H), 2.70 (t, J = 5.7 Hz, 2H), 2.38 (s, 6H), 2.33 (s, 6H), 2.13 (s, 3H), 2.09 (s, 3H). <sup>13</sup>C-NMR: (101 MHz, CDCl3) δ 157.14, 156.57, 150.46 (d, J = 245.5 Hz), 150.38 (d, J = 245.3 Hz), 146.37 (d, J = 10.8 Hz), 146.31 (d, J = 10.8 Hz), 141.40, 141.34, 139.51 (d, J = 4.2 Hz), 139.45 (d, J = 4.0 Hz), 137.55, 137.52, 134.77, 134.68, 134.52, 133.93, 131.91, 131.48, 131.19, 130.79, 130.57, 127.81, 127.22, 120.90 (d, J = 6.8 Hz), 120.81 (d, J = 6.7 Hz), 115.04 (d, J = 10.6 Hz), 114.86, 114.81, 114.60, 113.69, 113.25, 65.83, 65.68, 58.25, 58.18, 56.03 (d, J = 3.1 Hz), 56.01, 45.82, 45.76, 23.23, 23.02. MS (ESI): *m/z* = 440.30 [M+H]<sup>+</sup> (100%), *m/z* = 442.30 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.33 (94:6 methylene chloride: methanol).

*E/Z*-(2-{4-[1-(4-Chlorophenyl)-2-(4-fluoro-3-methoxyphenyl)-propenyl]-phenoxy} ethyl)-diethyl-amine (**28**)

C28H31ClFNO2. Yield: 52%. Orange oil. Purity: 97.68%. <sup>1</sup>H-NMR ((400 MHz, CDCl3) δ 7.31 (d, J = 7.0 Hz, 2H), 7.14 (dd, J = 16.5, 7.5 Hz, 4H), 7.02 (d, J = 7.2 Hz, 2H), 6.89 (t, J = 8.9 Hz, 4H), 6.81 (d, J = 7.5 Hz, 2H), 6.76 (d, J = 7.4 Hz, 2H), 6.73–6.65 (m, 3H), 6.65–6.57 (m, 3H), 4.17 (s, 2H), 4.04 (s, 2H), 3.63 (s, 6H), 2.97 (d, J = 30.6 Hz, 4H), 2.76 (d, J = 12.1 Hz, 8H), 2.13 (s, 3H), 2.10 (s, 3H), 1.19–1.08 (m, 12H). <sup>13</sup>C-NMR: (101 MHz, CDCl3) δ 158.81, 158.37, 151.95, 151.76, 145.82 (d, J = 9.7 Hz), 145.80 (d, J = 9.6 Hz), 142.03, 141.98, 140.95, 140.69, 138.17, 135.25, 134.70, 134.56, 133.47, 132.62, 132.14, 131.92, 131.46, 131.29, 128.49, 127.91, 121.36 (d, J = 6.6 Hz), 117.43 (d, J = 13.6 Hz), 115.26 (d, J = 13.6 Hz), 115.23, 114.34, 113.88, 56.20, 53.56, 53.38, 51.73, 51.59, 47.87, 47.83, 23.38, 23.20, 11.39, 11.31. MS (ESI): *m/z* = 468.30 [M+H]<sup>+</sup> (100%), *m/z* = 470.30 [M+H+2]<sup>+</sup> . R<sup>f</sup> : 0.50 (93:7 methylene chloride: methanol).

*3.2. Biology*

3.2.1. Yeast Estrogen Receptor Assay (YES)

The yeast estrogen receptor assay was supplied by Dr. J.P. Sumpter (Brunel University, Uxbridge, UK) and was used to determine the relative transactivation activity of the human ERα as formerly described [15]. Briefly, *Saccharomyces cerevisiae* stably transfected with a human ERα and an estrogen-responsive element fused to the reporter gene *lacZ* encoding for β-galactosidase were treated with the test substances for about 48 h. The β-galactosidase enzymatic activity was measured in a colorimetric assay using a microplate photometer by hydrolysis of the substrate chlorophenol red β-D-galactopyranoside (Roche Diagnostics, Mannheim, Germany), which leads to the formation of chlorophenol red. This can be measured as an increased absorption at 540 nm. All compounds were diluted in DMSO. 17β-estradiol (E2) (Sigma, Deissenhofen, Germany) 10 nM was used as a positive control, and DMSO was used as vehicle control. All compounds, also TAM (TAM) (Biotrend, Cologne, Germany), and 4-hydroxy-TAM (4-OH-TAM), were screened for agonistic and anti-estrogenic activity in a concentration of 1 µM; anti-estrogenic assays were performed in combination with 0.5 nM/1 nM E2 depending on the EC<sup>50</sup> value in each experimental series. All compounds were tested in technical quadruplicates and biological triplicates. Statistical analysis was performed by analysis of variance (ANOVA) and Tukey's post-hoc

test with the significance level of *p* < 0.05. The relative β-galactosidase activity of all compounds is shown in Tables 2–4.

#### 3.2.2. NCI Anti-Cancer Screening

All compounds were subjected to the NCI in vitro disease-oriented human cells screening panel assay. The human tumor cell lines of the cancer-screening panel are grown in RPMI 1640 medium containing 5% fetal bovine serum and 2 mM L-glutamine. For a typical screening experiment, cells are inoculated into 96-well microtiter plates in 100 µL at plating densities ranging from 5000 to 40,000 cells/well depending on the doubling time of individual cell lines. After cell inoculation, the microtiter plates are incubated at 37 ◦C, 5% CO2, 95% air, and 100% relative humidity for 24 h prior to the addition of experimental drugs. After 24 h, two plates of each cell line are fixed in situ with TCA to represent a measurement of the cell population for each cell line at the time of drug addition. Experimental drugs are solubilized in dimethyl sulfoxide at 400-fold the desired final maximum test concentration and stored frozen prior to use. At the time of drug addition, an aliquot of frozen concentrate is thawed and diluted to twice the desired final maximum test concentration with complete medium containing 50 µg/mL gentamicin. Additional four, 10-fold, or 1/2 log serial dilutions are made to provide a total of five drug concentrations plus control. Aliquots of 100 µL of these different drug dilutions are added to the appropriate microtiter wells already containing 100 µL of medium, resulting in the required final drug concentrations. Following drug addition, the plates are incubated for an additional 48 h at 37 ◦C, 5% CO2, 95% air, and 100% relative humidity. For adherent cells, the assay is terminated by the addition of cold TCA. Cells are fixed in situ by the gentle addition of 50 µL of cold 50% (*w/v*) TCA (final concentration, 10% TCA) and incubated for 60 min at 4 ◦C. The supernatant is discarded, and the plates are washed five times with tap water and air-dried. Sulforhodamine B (SRB) solution (100 µL) at 0.4% (*w/v*) in 1% acetic acid is added to each well, and plates are incubated for 10 min at room temperature. After staining, unbound dye is removed by washing five times with 1% acetic acid, and the plates are air-dried. Bound stain is subsequently solubilized with a 10 mM trizma base, and the absorbance is read on an automated plate reader at a wavelength of 515 nm. For suspension cells, the methodology is the same except that the assay is terminated by fixing settled cells at the bottom of the wells by gently adding 50 µL of 80% TCA (final concentration, 16% TCA). Compounds are screened at a dose of 10 µM, hits showing mean growth inhibition over 60 cell lines >50% are escalated for 5-dose screening assay. To construct a dose-response curve, about 60 cell lines of nine tumor subpanels were incubated with five concentrations (0.01–100 µM) for each compound. Three response parameters (GI50, TGI, and LC50) were calculated for each cell line. The GI<sup>50</sup> value corresponds to the compound's concentration causing a 50% decrease in net cell growth, the TGI value is the compound's concentration resulting in total growth inhibition, and the LC<sup>50</sup> value is the compound's concentration causing a net 50% loss of initial cells at the end of the incubation period (48 h) [41].

#### 3.2.3. Alkaline Phosphatase Activity in Ishikawa Cells

Estrogens stimulate the activity of alkaline phosphatase (AlkP) in Ishikawa cells (human endometrial adenocarcinoma cells; kindly provided by Prof. Masato Nishida, National Hospital Organization, Kasumigaura Medical Center, Japan). This enzyme activity is estimated by using the chromogen substrate (4-nitrophenylphosphate). These cells are very sensitive to estrogens; estradiol already induces the AlkP activity at a concentration of 10−<sup>12</sup> M [20]. The procedure was modified by Littlefield et al., 1990 [42].

Briefly, cells were cultured in DMEM/F12 medium without phenol red containing 5% dextran-coated charcoal-treated FCS (DCC, BioWest, (Nuaille, France) and insulin– transferrin–selenium A (Invitrogen, Karlsruhe, Germany). Cells were kept in plastic culture flasks at 5% CO<sup>2</sup> and 37 ◦C and harvested by brief exposure to trypsin (0.05%) EDTA at 37 ◦C. For experiments, the cells were seeded in 96-well plates at the required density

of 11,000 cells per well. Compounds, diluted in DMSO (Carl Roth GmbH, Germany), were tested in a concentration of 1 µM. DMSO was used as a negative control, 10 nM 17β-estradiol as a positive control, respectively. After 72 h incubation, cells were harvested, washed twice with PBS, and incubated at −80 ◦C for about 30 min to lyse the cells. After thawing, the lysates were resuspended in reaction buffer (274 mM mannitol, 100 mM CAPS, 4 mM MgCl2, pH 10.4) containing 4 mM p-nitrophenylphosphate (NPP). After incubation for 1 h in the dark, AlkP activity was assayed by using the hydrolysis of pnitrophenylphosphate to p-nitrophenol at pH 10.4 and the spectrometric determination of the kinetic of the product formation at 405 nm. All compounds were tested in technical triplicates and biological triplicates. Statistical analysis was performed by analysis of variance (ANOVA) and Tukey's post-hoc test with the significance level of *p* < 0.05.

#### 3.2.4. Uterotrophic Assay

The most common short-term in vivo assay for (anti)-estrogenicity is the uterine growth test, suitable for screening ERα agonists and antagonists. The primary endpoint is the uterine wet weight (UWW). An increase in UWW indicates an estrogenic activity of the test compound [43]. Sprague Dawley female rats (170–200 g) were obtained from the animal colony of the National Institute of Research (Cairo, Egypt). The rats were housed in a temperature-controlled room (23–24 ◦C) with a 12 h light:dark cycle and with free access to food and water. They were allowed to acclimatize to the animal house of the German University in Cairo for at least 1 week before initiating the experiments. All efforts were made to minimize animal discomfort and suffering. Animals were ovariectomized. After 14 days of endogenous hormonal decline, the animals were subcutaneously treated for three days with respective compounds. The animals were randomly allocated to treatment and vehicle groups (n = 6). 17β-estradiol were administrated s.c. at a dose of 10 µg/kg/d BW, all test compounds at a dose of 10 mg/kg/d BW daily for a period of three days. Animals were sacrificed by CO<sup>2</sup> inhalation after light anesthesia by inhaling an O2/CO<sup>2</sup> mixture around 24 h after the third administration. The uterus wet weight was determined.

#### *3.3. In Silico Study*

A docking experiment was implemented to dock compounds **3** into the active site of estrogen receptor α (ERα) with the program MOE version 2009.10. The Protein Data Bank (PDB) crystal structure of ERα co-crystallized with DES (3ERD) was imported into MOE [40]. All possible hydrogen atoms were added. Atomic charges were assigned using the MMFF94 force field parameters in MOE. The binding pocket was selected and extended 4.5 Å around the pocket. Compound **3E**, **3Z,** and DES were built using MOE builder; we run a conformational search to build a database (.mdb) of the most stable conformers of the three compounds. The.mdb file was then docked into the pocket, the poses from the ligand conformation were generated using alpha triangle, the scoring function used was London dG with no refinement. To ensure more accurate docking procedures, DES were redocked to the binding pocket using the same MOE settings as compound **3**. MOE was also used to represent the 2D interactions within ERα LBD.

#### **4. Conclusions**

Structural modifications on rings **A**, **B,** and **C** of TAM led to compounds with moderate to high estrogenic activity and potential growth inhibition activity on ER-positive and -negative breast cancer cell lines. Compounds **12** and **19** were tested in vivo in an ovariectomized rat model and are promising candidates for the development of novel SERMs with potent anti-neoplastic activity. This work opens the horizon for further development of triphenylethylenes where the para position of ring **C** bears different substituents; such structural modification can alter both their estrogenic/anti-estrogenic properties and have a prominent effect on their metabolic fate.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/ijms222212575/s1.

**Author Contributions:** Conceptualization, G.V., A.H.A. and N.S.A.; Methodology, H.E.E., M.M.R., N.O.A., M.A.V., O.Z. and N.S.A.; Supervision, J.J.W., K.S., J.W. and N.S.A.; Writing—original draft, J.W. and N.S.A.; Writing—review and editing, J.W., G.V., A.H.A. and N.S.A.; N.S.A. is the PI of the projects that partially financed the work, STDF grants #30298 and STDF grant #5386. N.S.A. is the coordinator of the ERA+ staff mobility grant. All authors have read and agreed to the published version of the manuscript.

**Funding:** This paper is based upon work supported financially by the Science and Technology Development Funding authority (STDF) under grant no.: 5386 and grant no.: 30298 to Nermin S. Ahmed.

**Institutional Review Board Statement:** The study was conducted according to the guidelines of the Declaration of Helsinki and approved by the Ethics Committee of German University in Cairo (March 2019). Animal procedures were performed following the approval of the Ethics Committee of the German University in Cairo in association with the recommendations of the National Institutes of Health Guide for Care and Use of Laboratory Animals (publication no. 85-23, revised 1985).

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** The authors are grateful to Susanne Broschk for her technical assistance in the biological screening. The authors are deeply grateful to the authority of the National Cancer Institute, USA, for the antitumor screening. Nouran O. Elbably thank the DAAD (German Academic Exchange Service) for an Erasmus+ stipend (code: **999897729**).

**Conflicts of Interest:** The authors declared that they have no conflict of interest.

#### **References**


### *Article* **Molecular Iodine/Cyclophosphamide Synergism on Chemoresistant Neuroblastoma Models**

**Winniberg Álvarez-León † , Irasema Mendieta † , Evangelina Delgado-González , Brenda Anguiano and Carmen Aceves \***

> Instituto de Neurobiología, Universidad Nacional Autónoma de México (UNAM) Juriquilla, Querétaro 76230, Mexico; winnsteph@gmail.com (W.Á.-L.); aliciairasema@hotmail.com (I.M.); edelgado@comunidad.unam.mx (E.D.-G.); anguianoo@unam.mx (B.A.)

**\*** Correspondence: caracev@unam.mx; Tel.: +52-442-238-1067

† W.Á.-L. and I.M. worked together and contributed equally to this paper.

**Abstract:** Neuroblastoma (Nb), the most common extracranial tumor in children, exhibited remarkable phenotypic diversity and heterogeneous clinical behavior. Tumors with MYCN overexpression have a worse prognosis. MYCN promotes tumor progression by inducing cell proliferation, de-differentiation, and dysregulated mitochondrial metabolism. Cyclophosphamide (CFF) at minimum effective oral doses (metronomic therapy) exerts beneficial actions on chemoresistant cancers. Molecular iodine (I<sup>2</sup> ) in coadministration with all-trans retinoic acid synergizes apoptosis and cell differentiation in Nb cells. This work analyzes the impact of I<sup>2</sup> and CFF on the viability (culture) and tumor progression (xenografts) of Nb chemoresistant SK-N-BE(2) cells. Results showed that both molecules induce dose-response antiproliferative effects, and I<sup>2</sup> increases the sensibility of Nb cells to CFF, triggering PPARγ expression and acting as a mitocan in mitochondrial metabolism. In vivo oral I2/metronomic CFF treatments showed significant inhibition in xenograft growth, decreasing proliferation (Survivin) and activating apoptosis signaling (P53, Bax/Bcl-2). In addition, I<sup>2</sup> decreased the expression of master markers of malignancy (MYCN, TrkB), vasculature remodeling, and increased differentiation signaling (PPARγ and TrkA). Furthermore, I<sup>2</sup> supplementation prevented loss of body weight and hemorrhagic cystitis secondary to CFF in nude mice. These results allow us to propose the I<sup>2</sup> supplement in metronomic CFF treatments to increase the effectiveness of chemotherapy and reduce side effects.

**Keywords:** neuroblastoma; molecular iodine; cyclophosphamide; xenografts; metronomic therapy

#### **1. Introduction**

Neuroblastoma (Nb) is the most common extracranial tumor in children accounting for 15% of pediatric oncology deaths. The overexpression of the neural MYC gene (MYCN) characterizes the chemoresistant Nb and has a worse prognosis [1]. MYCN induces cell proliferation, inhibits cell differentiation, and maintains the stem-like phenotype [2]. These levels correlate to metastasis and angiogenesis, and MYCN overexpression affects the mitochondria metabolism to support the higher energy demand of chemoresistant Nb cells [1,2]. One of the most widely used drugs in this pathology is cyclophosphamide (CFF), an effective and low-cost chemotherapy. CFF is a prodrug, biotransformed into two metabolites: phosphoramide mustard (amino-[bis(2-chloroethyl)amino] phosphinic acid), which is the active antineoplastic principle, and acrolein (prop-2-enal), associated with several side effects including inflammation and hemorrhagic cystitis [3]. The use of CFF in metronomic therapy has recently been proposed as an effective alternative for chemoresistant cancers [4]. Metronomic therapy uses chronic oral treatments with minimum effective doses that exert their effects by inhibiting angiogenesis, immune modulation of the tumor stroma, and apoptosis of tumor cells. Its long-term and low-dose use reduces side effects and maintains the patient's quality of life [4]. Furthermore, the use of combined therapies

**Citation:** Álvarez-León, W.; Mendieta, I.; Delgado-González, E.; Anguiano, B.; Aceves, C. Molecular Iodine/Cyclophosphamide Synergism on Chemoresistant Neuroblastoma Models. *Int. J. Mol. Sci.* **2021**, *22*, 8936. https://doi.org/ 10.3390/ijms22168936

Academic Editor: Angela Stefanachi

Received: 13 July 2021 Accepted: 16 August 2021 Published: 19 August 2021

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**Copyright:** © 2021 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 (https:// creativecommons.org/licenses/by/ 4.0/).

that include cellular re-differentiation or immune reactivation messengers are promising approaches that are currently being tested [5,6].

In addition, the antineoplastic effects of molecular iodine (I2) are well established, triggering apoptotic and redifferentiation mechanisms in several cancer cells, including mammary, ovary, and prostate, among others [7]. These effects are mediated partially by the activation of peroxisome proliferator-activated receptors type gamma (PPARγ) and directly, as a mitocan element, by thiol depletion and disruption of the mitochondrial membrane potential (Mmp), triggering the intrinsic apoptosis [8]. Furthermore, our previous report showed that the I<sup>2</sup> supplement sensitized Nb cells to all-trans retinoic acid (ATRA) in vitro and synergized the antitumor effect of ATRA preventing body-weight loss and diarrhea episodes in nude mice [9]. The present work analyzes the impact of I<sup>2</sup> and metronomic CFF supplements on the viability (culture) and tumor progression (xenografts) of chemoresistant neuroblastoma SK-N-BE(2) cells [10].

#### **2. Results**

*2.1. Results In Vitro*

2.1.1. Viability

Figure 1A shows the effect of molecular iodine (I2), CFF, and their combination on the viability of the SH-SY5Y and SK-N-BE(2) cell lines at 96 h. Both components generate similar dose-response action, being SH-SY5Y the most sensitive and corroborating that SK-N-BE(2) is a highly resistant cell line. I<sup>2</sup> supplementation showed an IC50 of 205.5 µM in SH-SY5Y cells and 343.7 µM in SK-N-BE(2) cells (Figure S1). In CFF, the IC50 was 0.602 µM in SH-SY5Y and 1.045 µM in SK-N-BE(2) (Figure S2). Specific time-response data are summarized in Figures S1 and S2. All further experiments were analyzed in SK-N-BE(2) cells. The combination of 200 µM I<sup>2</sup> with two concentrations of CFF confirmed that the presence of iodine increases the sensibility of these cells to CFF, improving response by 15% at 1.0 µM CFF and up to 40% at 0.5 µM CFF (Figure 1B). The combination index (CI) for I2/CFF treatments presented a CI value of 0.00835 at 0.5 µM CFF and 3.39 at 1 µM CFF, indicating synergism at 0.5 µM CFF. To analyze the participation of PPARγ in the I<sup>2</sup> response, we used the agonist rosiglitazone (RGZ 5.0 µM) in the presence or absence of the antagonist GW9662 (1.0 µM) for 96 h. RGZ showed a similar inhibitory effect in viability observed with I<sup>2</sup> (34 vs. 28%). The preincubation (2 h before) with GW9662 canceled the RGZ inhibitor effect but had a partial impact on the I<sup>2</sup> supplement (24%), suggesting that I<sup>2</sup> exerts its actions through other mechanisms besides PPARγ (Figure 1C).

#### 2.1.2. Apoptosis

Figure 2 shows the percentage of apoptosis-positive cells (annexin Cy5) at two different times. At 48 h, 200 µM I<sup>2</sup> and 1.0 µM CFF exhibited similar apoptotic induction (~35% each), and the adjuvant action (I<sup>2</sup> + CFF group) enhanced the apoptosis effect to 40%. At 96 h, apoptosis was maintained primarily in I<sup>2</sup> groups. The apoptotic index Bax/Bcl-2 (RT-PCR), an indicator of caspase pathway activation, indicated that both components exert significant induction, showing a predominant action of I<sup>2</sup> at both times.

#### 2.1.3. Mitochondrial Activity

Figure 3 depicts the fluorescence generated by the MitoTracker entrance after I<sup>2</sup> and CFF treatments. Iodine exerted a rapid and significant increment in mitochondrial permeability at 12 h and maintained until 48 h. In contrast, CFF induced a change in mitochondria metabolism after 48 h. The presence of both components exhibited a synergistic effect only at 48 h.

**Figure 1.** Effect of I<sup>2</sup> , CFF and their combination on the viability (exclusion dye trypan blue) of neuroblastoma cell lines. (**A**) Dose response of I<sup>2</sup> and CFF in SH-SY5Y and SK-N-BE(2) cells. (**B**) Synergistic effect (CI < 1.0) of 200 µM I<sup>2</sup> with 0.5 µM CFF (reduction 45%) and 1.0 µM CFF (reduction 25%) concentrations in SK-N-BE(2) cells. (**C**) Participation of PPARγ in the effect of I<sup>2</sup> analyzed with rosiglitazone (RGZ; PPARγ agonist) and GW9662 (PPARγ antagonist) in SK-N-BE(2) cells. All experiments were carried out for 96 h. Data are representative of three independent experiments per triplicate and are expressed as the mean ± SD. Different letters denote statistical differences per group (*p* < 0.05).

#### 2.1.4. Molecular Response

Figure 4 shows the effect of treatments on gene expression. I<sup>2</sup> supplements increased the genes associated with differentiation (PPARγ) and diminished those associated with aggressiveness (MYCN) and resistance (Survivin; SVV). The presence of CFF only reduced SVV expression. In the I<sup>2</sup> + CFF group, only the effects of I<sup>2</sup> remained, suggesting that I<sup>2</sup> is the inductor of differentiation. The multidrug resistance gene MDR1 did not exhibit differences with any treatment.

**Figure 2.** Apoptotic induction by I<sup>2</sup> and CFF supplementation in the neuroblastoma SK-N-BE(2) cells. The cell line was supplemented with 200 µM I<sup>2</sup> , 1 µM CFF, and I2/CFF. (**A**) Apoptotic-positive cell percentage was evaluated with the Attune flow cytometer at 48 and 96 h; representative dot plots at 48 h and quantitative results are shown; (**B**) the apoptotic Bax/Bcl-2 index was assessed using real-time PCR at 48 and 96 h. Figures are representative of three independent experiments per triplicate. Data are expressed as the mean ± SD. Different letters denote statistical differences (*p* < 0.05).

**Figure 3.** *Cont.*

**Figure 3.** Effect of I<sup>2</sup> and CFF on the mitochondrial functional state in SK-N-BE(2) cells. The cell line was supplemented with 200 µM I<sup>2</sup> , 1 µM CFF, and I<sup>2</sup> + CFF, for 12, 24, and 48 h to evaluate the mitochondrial permeability (20 µM, scale bar). Representative micrographs (48 h) and bar graph for all times of the MitoTracker signal (Mmp ψ: mitochondria membrane potential change; RFU: Relative Fluorescence Units). Figures are representative of three independent experiments per triplicate. Data are expressed as the mean ± SD. Different letters denote statistical differences (*p* < 0.05).

**Figure 4.** Effect of I<sup>2</sup> and CFF on gene expression in the neuroblastoma SK-N-BE(2) cell line. I<sup>2</sup> (200 µM), CFF (1 µM), and both (I<sup>2</sup> + CFF) were supplemented for 96 h. Aggressive [neural MYC (MYCN)] and

chemoresistant markers (Survivin (SVV), and multidrug resistance mutation 1 gene (MDR1)) were analyzed by RT-qPCR. PPARγ expression and protein content was analyzed by RT-qPCR (PPARγ/βactin) and Western blot (PPARγ/Actin). Figures are representative of three independent experiments per triplicate. Data are expressed as the mean ± SD. Different letters denote statistical differences (*p* < 0.05).

#### *2.2. Results In Vivo*

#### 2.2.1. Tumor Growth

Nude mice with SK-N-BE(2) xenografts were used to analyze the effects of I<sup>2</sup> and CFF under in vivo conditions. Treatments were given when the tumor reached a size of 1.5 cm<sup>3</sup> . We evaluated the impact of oral metronomic CFF doses (20 mg/kg/day; 0.06%) with or without the I<sup>2</sup> supplement (8 mg/kg/day; 0.025%) for three weeks. Previous studies from our laboratory found that the presence of xenografts generates a small but consistent reduction in body weight gain (BWG) in all animals and that the I<sup>2</sup> supplement prevents this loss. Figure 5 shows that I2-supplemented animals exhibited an increase in BWG like that of the control group of xenograft-free (x.f.) animals. The animals with tumors, both the control and the CFF groups, showed a reduction in BWG from the first week. In contrast, the combined group (I<sup>2</sup> + CFF) recovered BWG in the last week of treatment, indicating a beneficial effect of I2.

**Figure 5.** Effect of I<sup>2</sup> and CFF on nude mice and xenografts. Nude mice with SK-N-BE(2) xenografts were supplemented with I<sup>2</sup> (0.025%) and metronomic CFF (0.060%) in drinking water for three weeks. The line graphs show the body weight gain, % tumor growth and final tumor volume compared to the control group. The pictures are representative of the tumor bleeding appearance. Data are expressed as the mean ± SD (*n* = 4). Different letters and \*\* denote statistical differences for the control group (*p* < 0.01).

Concerning tumor progression, after the third week, all treatments significantly inhibited tumor growth. The tumor volume decreased 44.72% with the I<sup>2</sup> supplement and 68.11% with CFF. The coadministration of both components (I<sup>2</sup> + CFF) showed a considerable, yet not statistically significant, tumor inhibition, decreasing the tumor size by 78.78%.

A characteristic of these types of tumors is their aberrant vascularization and abundant bleeding, and so they are known as blue tumors. As shown in Figure 5, I<sup>2</sup> supplementation considerably decreased bleeding patterns regardless of tumor size. However, the CFF group alone showed a decrease in tumor size but did not change bleeding appearance.

#### 2.2.2. Histopathology

Figure 6 shows the analysis of vascularization (H&E stain and CD34 immunohistology and quantification) and collagen fibrosis content (Masson's stain and quantification). Control and CFF xenograft micrography showed aberrant vascular patterns and abundant extravascular erythrocytes (H&E stain). In contrast, treatments with I<sup>2</sup> alone and combined (I<sup>2</sup> + CFF) showed a consistent reduction in the vasculature, minor invasion of tumor cells in vascular structures, and fewer extravascular erythrocytes. The quantification of mean vascular density (MVD) by CD34 immunohistochemistry showed that I<sup>2</sup> and CFF decrease vasculature area in comparison with control group. A significant increase in positive collagen fibers (blue stain) is observed in I<sup>2</sup> and CFF groups suggesting a hypoxic microenvironment, with tumoral cell death fibrosis substitution. Following this hypoxic status, I2-treated tumors exhibited an elevated expression of Hypoxia-inducible factor (HIF1) but not vascular endothelial growth factor (VEGF), which corroborates that the I<sup>2</sup> treatments do not induce an increase in vascularization.

**Figure 6.** Effect of I<sup>2</sup> and CFF on the histopathology of SK-N-BE(2) xenografts. H&E stain (40×). Immunohistochemistry of endothelial protein CD34 and quantification of mean vascular density (area CD34+/field). Masson's trichrome stain and percent of positive fibrosis area. Epithelial (red) and collagen fibers (blue) (40×) (50 µM, bar graph). Hypoxia-inducible factor (HIF1) and vascular endothelial growth factor (VEGF) expression (RT-qPCR). Data are expressed as the mean ± SD (*n* = 4). Different letters denote statistical differences (*p* < 0.05).

#### 2.2.3. Molecular Response

Similar to the in vitro results, xenografts from the control animal expressed an elevated amount of MYCN (Figure 7). The I<sup>2</sup> supplement modified the xenografts' aggressivity pattern, increasing the expression of differentiation promoters (PPARγ and TrkA) and decreasing those related to resistance (MYCN and TrkB). Interestingly, the higher TrkB expression promoted by the CFF supplement was suppressed by the presence of I<sup>2</sup> (I<sup>2</sup> + CFF group). In addition, both components (I<sup>2</sup> and CFF) induced the expression of p53 and increased the apoptotic index Bax/Bcl-2.

**Figure 7.** Effect of I<sup>2</sup> and CFF on gene expression in neuroblastoma xenografts. Nude mice with SK-N-BE(2) xenografts were supplemented with I<sup>2</sup> (0.025%) or the metronomic dose of CFF (0.060%) administered in drinking water for three weeks. Genes related to neuronal differentiation (PPARγ and TrkA), aggressiveness (MYCN and TrkB), and apoptosis induction (p53 and Bax/Bcl-2 index) were analyzed by RT-qPCR. PPARγ content was analyzed by Western blot. Data are expressed as the mean ± SD (*n* = 4). Different letters denote statistical differences (*p* < 0.05).

#### 2.2.4. Preventive Effect of I<sup>2</sup> in Bladder Damage

The more frequent side effect of CFF treatment is hemorrhagic cystitis, evidenced by hypervascularity, edema, inflammation, and bleeding. We examined the bladder morphology at the end of the experiment and evaluated the vasculature (blood vases/field) through expression of CD34 (immunohistochemistry) and histopathology (H&E) (Figure 8). No significant differences in the vasculature were found in the treatments. However, the stains with H&E revealed clear signs of edema in the lamina propria and an increased thickness in the urothelium in the CFF group (arrows). I<sup>2</sup> and I<sup>2</sup> + CFF group bladders did not show these alterations, indicating the I<sup>2</sup> alone did not cause any irritation and that, in the presence of CFF, this halogen exerts a significant preventive action.

**Figure 8.** Effect of I<sup>2</sup> and CFF on the bladder morphology of the chemoresistant neuroblastoma model. The figure shows the urothelial thickness indicated by arrows in H&E representative micrographs (100 µM, bar graph) and the vasculature quantification (blood vessel number/field) of CD34 immunohistochemical staining (representative micrographs and bar graph). Three sections per tumor sample were analyzed. Data are expressed as the mean ± SD (*n* = 4). Different letters denote statistical differences (*p* < 0.05).

#### **3. Discussion**

Previously our group showed that I<sup>2</sup> in coadministration with ATRA synergizes apoptosis and cell differentiation in Nb cells [9]. The present work analyzes the impact of I<sup>2</sup> and CFF in viability and tumor progression of Nb chemoresistant SK-N-BE(2) cells. Our results corroborate that these cells have a high resistance to various antineoplastic components since both I<sup>2</sup> and CFF exert attenuated effects up to 40% compared to the more sensitive SH-SY5Y cells [10]. However, an important finding is that the I<sup>2</sup> supplement, which does not generate any side effects, increased the sensitivity to CFF by 25 to 40%. It is well established that the primary mechanism of CFF is the induction of p53 apoptosis via DNA adducts formation [11]. Conversely, I<sup>2</sup> actions are more complex since this halogen could act directly on mitochondria by inducing an apoptosis cascade [12], or indirectly by activating PPARγ and triggering redifferentiation or apoptotic signaling [7,13]. Our results showed that both components (I<sup>2</sup> and CFF) increased apoptosis (exposure of annexin-Cy5 and low expression of SVV), but only I<sup>2</sup> groups modified master differentiation genes, decreasing the expression of MYCN, and significantly inducing PPARγ. Iodine in neoplastic cells binds with arachidonic acid and generates an iodolipid called 6-iodolactone (6-IL) [14,15]. This iodolipid is a specific activator of PPARγ [16,17]. These receptors are

expressed in Nb, and the use of agonists impairs proliferation and induces differentiation of these cells [9,18]. PPARγ agonists reduce levels of MYCN by inhibiting critical molecules within the PI3K/AKT/mTOR signaling pathway increasing GSK-3β activity as well as MYCN phosphorylation and its proteasome degradation [19–21]. We corroborated that PPARγ reduces the viability of these SK-N-BE(2) Nb cells since the supplement with 1 uM RGZ decreased its proliferation. Moreover, we found that the I<sup>2</sup> effect was partially canceled with the agonist GW9662, suggesting that the antineoplastic effects of I<sup>2</sup> include the activation of PPARγ, but other mechanisms also contribute.

Mitochondria metabolism is considered a hallmark of cancer, showing a crucial contributor in the process as metabolic reprogramming, generation of reactive oxygen species (ROS) and production of metabolites that enhance oncogenesis [22]. Recently, several therapeutic approaches for these processes identified the "mitocans," a category of drug that targets the mitochondria of cancer cells [8]. Many natural agents can target mitochondria and exert anticancer activities with minimal or no side effects. In the light of this process, I<sup>2</sup> seems to be a mitocan since, in cancer cells, I<sup>2</sup> depleted thiol generation and disrupted the Mmp, inducing significant increases in its permeability and triggering apoptosis [12,23]. The substantial rise in MitoTracker signaling observed in I<sup>2</sup> groups starting at 12 h corroborated these direct mitochondrial effects and explains, in part, the increase in CFF sensibility. Previous data from our laboratory showed that this increase in Mmp was accompanied by a decrease in SVV content in both intramitochondrial and cytoplasmic compartments [9]. SVV is an apoptosis-inhibiting factor (IAP) [24]. SVV overexpression in Nb cells makes them resistant to ATRA, protecting them against agents that damage DNA, and stabilizes the mitochondrial membrane by decreasing apoptosis induction [20,25].

The xenograft model was efficient and reproducible since we obtained between 95 and 100% implantation. Moreover, we corroborated that this is an aggressive Nb type generating fast-growing tumors and hypervascularization characteristics (bluish). The results showed that both the metronomic CFF and the I<sup>2</sup> supplement effectively decreased tumor growth (68.11% and 44.72%, respectively), especially when both components were co-administered (78.78%). In addition, the I<sup>2</sup> supplement was accompanied by decreased aberrant vascularization and bleeding, associated with a significant increase in the expression of HIF1, which is the first signaling secondary to lack of oxygen. No change in the expression of VEGF, the inducer of new vessels, was observed. This pattern can be interpreted as a modification in the vascular cytoarchitecture rather than a process of angiogenesis, but the possible mechanisms involved in this I<sup>2</sup> effect have not yet been elucidated. However, conventional mechanisms of I<sup>2</sup> could participate. Angiogenesis is an active process that involves a significant increase in ROS generation, promoting the angiogenic switch from quiescent to active endothelial cells [25,26]. It is possible that the mitocan effect of iodine neutralized ROS [8]. An alternative way might be the significant decrease in MYCN expression observed with I<sup>2</sup> treatments. MYCN induces angiogenesis by its direct action in VEGF amplification [2].

At molecular level, the decrease in tumor size was accompanied by an increase in the apoptosis markers p53 and Bax/Bcl-2 index in all treated groups. This apoptotic induction also agrees with the finding that the tumors supplemented with both components had a higher proportion of type 1A collagen fibers (Masson's trichrome staining), indicating the replacement of epithelial cancer cells with fibrous tissue in response to rapid induction of apoptosis.

In addition, one important finding is the differential gene response between tumors treated with I<sup>2</sup> vs. CFF. Our results agreed with the in vitro results, that only I<sup>2</sup> supplements exert an evident modulation in the differentiation master genes increasing PPARγ and TrkA, decreasing the basal expression of MYCN and TrkB, and canceling the increase of TrkB secondary to CFF treatment. MYCN amplification in Nb is typically associated with epigenetic abnormalities to impair apoptosis, followed by the overexpression of the anti-apoptotic proteins Bcl-2, SVV, and TrkB [1,19,20]. TrkB stimulates cell survival and angiogenesis and activates the survival pathway PI3K/AKT, contributing to increased drug resistance [26,27]. Therefore, the rise in TrkB in the presence of CFF could be interpreted as a response of tumor cells to the drug's toxic effect. Studies analyzing this hypothesis are needed; however, the prevalence of I<sup>2</sup> action indicates an antitumor benefit.

Finally, we explored the possible role of the I<sup>2</sup> supplement in the side effects prevention associated with xenograft signalization and the bladder injury secondary to CFF administration. Previous studies in our group had detected that xenografts and tumor growth generate stress in the mouse, evidenced in the loss of BWG [9]. This effect has been described in preclinical and clinical studies and is known as cachexia [28]. Cachexia is accompanied by loss of adipocytes and muscle tissue with chronic inflammation and increases in proinflammatory factors such as TNFα and IL-6 [28,29]. The prevention of weight loss in the I2-supplemented groups might be due to two conditions: first, the antineoplastic effect of I<sup>2</sup> that prevents tumor growth and decreases the tumor mass signaling; and second, a direct impact on chronic inflammation processes due to I<sup>2</sup> antioxidant action [9,30]. This effect also appears to be exercised in the prevention of bladder injury. It is well known that the hepatic biotransformation of CFF produces, in addition to phosphoramide mustard (an antineoplastic metabolite), acrolein, which causes hemorrhagic cystitis [3]. We did not expect severe effects of acrolein at the metronomic dose used; however, the histological evaluation showed a thickening of the urothelium and the lamina propria of the bladder, which indicates moderate hemorrhagic cystitis. The I<sup>2</sup> supplement in coadministration with CFF prevented these alterations. This protective mechanism might be due to its antioxidant action. Previously, it has been shown that the chemical form of I<sup>2</sup> has an in vitro reducing capacity (FRAP test) ten times greater than ascorbic acid and 60 times greater than potassium iodide [31]. In vivo studies showed that the iodine supplement decreases the oxidative potential in the serum of rodents and patients [30]. The administration of other antioxidants, such as ascorbic acid, retinol, and resveratrol, improves oxidative stress by reducing ROS levels in bladder tissues generated by CFF treatment [32,33]. An unexplored alternative is that the I<sup>2</sup> could be binding directly to acrolein, decreasing its irritating action, and preventing its contact or entry into the urothelium of the bladder. This alternative is based on the acrolein structure that contains double bonds capable of being iodinated [34].

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

#### *4.1. Chemicals and Reagents*

The Cyclophosphamide for in vivo assays and the CFF active metabolite 4-Hydroperoxycyclophosphamide for in vitro assays were obtained by Cryofarma (Jalisco, Mexico) and Toronto Research Chemicals (Toronto, Ontario, CA, USA) respectively, and we denominate both with the same abbreviation (CFF). Rosiglitazone (RGZ; PPARγ-specific agonist, by Cayman Chemical, Los Angeles, CA, USA), GW9662 (PPARγ-specific antagonist, by Corning, Bedford, MA, USA) and Matrigel (basement membrane matrix, Corning, Bedford, MA, USA). Sublimed iodine was obtained from Macron-Avantor (Center Valley, PA, USA). The concentration of iodine solutions was verified by sodium thiosulfate titration. All other chemicals were of the highest purity grade available.

#### *4.2. Cell Culture*

The Nb cell lines SH-SY5Y (CRL-2266) and SK-N-BE(2) (CRL-2271) were obtained from the company American Type Culture Collection (ATCC, Manassas, VA, USA). All the experiments were performed with passages 1–5 and recently tested and authenticated by STR profiling (BIMODI Invoice number 190320-029). The conditions for cell culture were Dulbecco's Modified Eagle's Medium (DMEM) supplemented with fetal bovine serum (FBS, 10%) and penicillin/streptomycin (2%) by Invitrogen (Carlsbad, CA, USA) in a humidified chamber with 5% CO<sup>2</sup> atmosphere and 95% air at 37 ◦C.

#### *4.3. Cell Viability*

A total of 50,000 cells/well were seeded onto 12-well plates. After 24 h, different concentrations of CFF (0.5, 1, and 2 µM), I<sup>2</sup> (100, 200 and 400 µM) and I<sup>2</sup> + CFF (200 + 0.5 or 200 + 1.0, respectively) were added for 0, 24, 48, 72, and 96 h. Control groups were followed at the same times using deionized water as treatment (vehicle of I<sup>2</sup> and CFF). In the GW9662/RGZ or I2-treated groups, GW9662 (1 µM) was administered 2 h before RGZ (5 µM) or I<sup>2</sup> (200 µM) treatment.

After treatment, cells were detached and mixed with the exclusion dye trypan blue (0.04%) to count the cells using a hemocytometer in light microscopy; viability was reported as fold change against control. All experiments were carried out in three independent experiments per triplicate. To measure the extent of interaction between I<sup>2</sup> and CFF, data were analyzed by CompuSyn software 1.0 (ComboSyn, Inc., Paramus, NJ, USA) based on the combination index (CI) of the multiple drug effect equation of Chou-Talalay [35].

#### *4.4. Apoptosis*

Apoptosis was evaluated by flow cytometry using the apoptosis kit (ABCAM No. 14190, Cambridge, UK) with the Attune NXT flow cytometer (BRVY). Briefly, the pellet of cells was resuspended in PBS, and the monoclonal antibody for annexin and propidium iodide were added, according to manufacturer's instructions. The mixture was incubated for 30 min at room temperature and protected from light. After incubation, the cells were washed twice with PBS and resuspended in 500 µL PBS. Data analysis of 10,000 events was performed using FlowJo v10 (Trial version) software.

#### *4.5. Gene Expression*

TrkA, TrkB, PPARγ, SVV, MDR-1, MYCN, P53, Bax, Bcl-2, VEGF, HIF1, and β-actin were analyzed by RT-qPCR from SK-N-BE(2) cell cultures and xenografts after the corresponding treatments. Briefly, total RNA was obtained using Trizol reagent (Life Technologies, Inc., Carlsbad, CA, USA). RNA (2 µg) was reverse transcribed (RT) using oligodeoxythymidine (Invitrogen, Waltham, MA, USA). Real-time PCR was performed on the Rotor-Gene 3000 sequence detector system (Corbett Research, Mortlake, NSW, Australia) using SYBR Green as a DNA amplification marker (gene-specific primers are listed in Table 1). Relative mRNA levels were normalized to the mRNA level of β-actin.


**Table 1.** Oligonucleotide sequences.

#### *4.6. Mitochondrial Membrane Potential*

After 12, 24, and 48-h treatments (I2, CFF, or I<sup>2</sup> + CFF), the cells were PBS-washed and labeled with 200 nM MitoTracker Red CM-H2Xros (Thermo Fisher; Waltham, MA, USA) for 45 min. Then, cells were fixed for 10 min with ethanol, PBS-washed, and mounted with anti-FADE and DAPI. Micrographs were taken with an epifluorescence microscope (Axio Imager, Carl Zeiss, Jena, Germany). The software Image J 1.8 (National Institutes of Health, Bethesda, MD, USA) was used to quantify the relative fluorescence units (RFUs) and determine the mitochondrial functional state.

#### *4.7. Tumoral Implantation and Progression*

Xenografts with SK-N-BE(2) cells were generated using 5 × 10<sup>6</sup> cells/injection of Nb cells in 6–7-week-old male immunodeficient athymic nude mice (Foxn1 nu/nu, Harlan Mexico, Ciudad de Mexico, Mexico) as previously described [36]. Mice were housed in barrier conditions under a 12-h light/dark cycle with food and water supplied ad libitum. All the procedures followed the Animal Care and Use Program of the National Institutes of Health (NIH; Bethesda, MD, USA) and were approved by the Ethics Committee of the Instituto de Neurobiología (ethical approval number 035).

When palpable tumors reached a volume of 1 cm<sup>3</sup> , animals were randomly assigned to each group (*n* = 4). I<sup>2</sup> (8 mg/kg/day; 0.025%), CFF (20 mg/Kg/day; 0.060%), or a mixture of both. The treatments were supplied in drinking water ad libitum. The control group received only water. Animals were sacrificed after anesthesia with a ketamine/xylazine mixture (30 mg/Kg and 6 mg/Kg from Pisa Agropecuaria, Hgo., Mexico, and Cheminova CDMX, Mexico, respectively). The bladder and a tumor section were fixed in 10% formalin for at least 24 h and processed for immunohistochemistry. The remaining tumors were frozen in dry ice for RNA analysis.

#### *4.8. Immunohistochemistry*

Tumor sections and bladders were stained with hematoxylin-eosin (H&E) and Masson's trichrome techniques for histopathological analysis. In addition to the vasculature analysis, the endothelial protein antibody CD34 (ab182981; 1:2500, Abcam, Cambridge, UK) was used to detect endothelial-positive cells (Vector Labs, Burlingame, CA, USA). Sections were counterstained by hematoxylin. The Mean Vascular Density (CD34/mm<sup>2</sup> ) or vascular number per field were quantified by randomly analyzing three fields from three different sections of each tumor and bladder, using the software ImageJ version 1.8 (National Institutes of Health, Bethesda, MD, USA).

#### *4.9. Western Blot*

Western blot (WB) analysis of PPARγ proteins for tumor tissue was performed with the chemiluminescence technique [9]. Briefly, 50 µg of protein per lane were separated by electrophoresis in 10% acrylamide gel, proteins were later transferred to a nitrocellulose membrane (Bio-Rad, Hercules, CA, USA). The unspecified reaction was blocked overnight with PBS containing 5% skimmed milk powder. The membranes were treated with polyclonal antibodies (Santa Cruz Biotechnology, Los Angeles, CA, USA) against anti-PPARγ (ab209350, 54 kDa, 1: 1000, Abcam, Cambridge, MA, USA). As a secondary antibody, goat anti-rabbit (Thermo scientific 656120, 1: 10,000, Invitrogen, Waltham, MA, USA) was used. Proteins were visualized using chemiluminescent detection (ECL, Amersham Biosciences, Buckinghamshire, UK). The blots were visualized and pictured with Image LabTM (Biorad), and the densitometry analysis was performed with Image ImageJ V1.53e; PPAR levels were normalized to total protein of Ponceau red staining signal [37].

#### *4.10. Statistical Analysis*

Data for in vitro experiments are the media of three independent tests in triplicate. In vivo, four animals per group were used. Tissue analysis for PCR is the average of four samples, and three sections of each tumor were used for immunohistochemistry. Statistical analysis was performed by one-way ANOVA followed by Tukey's test for analysis between groups. Values with *p* < 0.05 were considered statistically significant.

#### **5. Conclusions**

Molecular iodine exerts antiproliferative and differentiation effects in Nb cell lines, increasing their sensitivity to CFF. Molecular mechanisms include decreased expression of master regulators related to malignancy (MYCN, TrkB), remodeling of the vasculature, and increased differentiation signaling (PPARγ and TrkA). Furthermore, I<sup>2</sup> supplementation prevents loss of body weight and hemorrhagic cystitis secondary to CFF in nude mice. These results allow us to propose the I<sup>2</sup> supplement in metronomic CFF treatments to increase the effectiveness of chemotherapy and reduce side effects.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10.339 0/ijms22168936/s1.

**Author Contributions:** W.Á.-L. and I.M. contributed equally to this paper. W.Á.-L. and I.M. performed the SK-N-BE(2) in vitro and in vivo assays and prepared the original draft. E.D.-G. supervised and analyzed the real-time polymerase chain reactions (RT-qPCR). B.A. provided coordination, supervised the statistical analysis, and corrected the draft. C.A. conceived the study, coordinated the research, and corrected the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was partially funded by PAPIIT-UNAM, grant numbers 203919, 205920; DGAPA postdoctoral fellowship award for Irasema Mendieta and CONACYT fellowship for Winniberg Álvarez-León.

**Institutional Review Board Statement:** The study was conducted according to the Animal Care and Use Program of the National Institutes of Health (NIH; Bethesda, MD, USA) and approved by the Research Ethics Committee of the Instituto de Neurobiología (ethical permit number 035).

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author.

**Acknowledgments:** The authors are grateful to Laura Ines Garcia, Elsa Nydia Hernández Ríos, Ericka de los Rios, Martín García Servín, Alejandra Castilla, and Antonieta Carbajo from INB-UNAM; Alejandra Castillo Carbajal, Carina Uribe Díaz and Rafael Palacios de la Lama from LIIGH-UNAM for technical assistance; Francisco Javier Valles and Rafael Silva for bibliographic assistance; Nuri Aranda and Sofia Gutierrez Ramirez for academic support; Alberto Lara, Omar Gonzalez, Ramon Martinez, and Maria Eugenia Rosas Alatorre for computer assistance; and Jessica Gonzalez Norris for proofreading.

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

#### **References**


### *Review* **Strategies to Improve the Antitumor Effect of** γδ **T Cell Immunotherapy for Clinical Application**

**Masatsugu Miyashita 1,2, \*, Teruki Shimizu 1 , Eishi Ashihara <sup>3</sup> and Osamu Ukimura 1**


Kyoto 607-8414, Japan; ash@mb.kyoto-phu.ac.jp

**\*** Correspondence: miyash@koto.kpu-m.ac.jp; Tel.: +81-752515595

**Abstract:** Human γδ T cells show potent cytotoxicity against various types of cancer cells in a major histocompatibility complex unrestricted manner. Phosphoantigens and nitrogen-containing bisphosphonates (N-bis) stimulate γδ T cells via interaction between the γδ T cell receptor (TCR) and butyrophilin subfamily 3 member A1 (BTN3A1) expressed on target cells. γδ T cell immunotherapy is classified as either in vivo or ex vivo according to the method of activation. Immunotherapy with activated γδ T cells is well tolerated; however, the clinical benefits are unsatisfactory. Therefore, the antitumor effects need to be increased. Administration of γδ T cells into local cavities might improve antitumor effects by increasing the effector-to-target cell ratio. Some anticancer and molecularly targeted agents increase the cytotoxicity of γδ T cells via mechanisms involving natural killer group 2 member D (NKG2D)-mediated recognition of target cells. Both the tumor microenvironment and cancer stem cells exert immunosuppressive effects via mechanisms that include inhibitory immune checkpoint molecules. Therefore, co-immunotherapy with γδ T cells plus immune checkpoint inhibitors is a strategy that may improve cytotoxicity. The use of a bispecific antibody and chimeric antigen receptor might be effective to overcome current therapeutic limitations. Such strategies should be tested in a clinical research setting.

**Keywords:** γδ T cells; immunotherapy; tumor resistance; combination therapy; tumor microenvironment; immune checkpoint inhibitor

#### **1. Introduction**

Cancer is one of the most serious and potentially fatal diseases in humans. According to statistical reports, there were an estimated 18.1 million new cancer cases and 9.6 million cancer-related deaths worldwide in 2018 [1]. Surgery, chemotherapy, and radiotherapy are the three pillars of antitumor therapy. Surgery and radiotherapy are curative for localized cancers; however, most cancer-related deaths are due to metastasis, which requires systemic therapy. Chemotherapy is the first-line systemic therapy against metastatic cancers; however, many cancers become resistant, which leads to treatment failure. Recently, immunotherapy, now regarded as the fourth pillar of antitumor therapy, has been used for systemic antitumor therapy.

T cell-based immunotherapy is an effective cancer treatment strategy. T cells are divided into two major subpopulations based on surface expression of αβ and γδ T cell receptors (TCRs). αβ T cells recognize peptide antigens in the context of non-self; for example, antigens expressed by cancer cells. αβ T cells are effector cells that operate within the adaptive arm of the immune system; these cells exert cytotoxicity in a major histocompatibility complex (MHC)-restricted manner. However, due to loss of MHC molecules, tumor cells are often resistant to attack by αβ T cells [2]. By contrast, γδ T cells are effectors that operate within the innate arm of the immune system; these cells act in an

**Citation:** Miyashita, M.; Shimizu, T.; Ashihara, E.; Ukimura, O. Strategies to Improve the Antitumor Effect of γδ T Cell Immunotherapy for Clinical Application. *Int. J. Mol. Sci.* **2021**, *22*, 8910. https://doi.org/ 10.3390/ijms22168910

Academic Editor: Angela Stefanachi

Received: 14 July 2021 Accepted: 17 August 2021 Published: 18 August 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 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 (https:// creativecommons.org/licenses/by/ 4.0/).

MHC-unrestricted manner, making them interesting mediators of cancer immunotherapy. Human γδ T cells were first identified in the mid-1980s [3–5]. They are abundant in the intestine and skin and play a role in defense against microbial infections in an MHCunrestricted manner [6]. Recent studies show that γδ T cells exert potent cytotoxic effects against various types of cancer cell [7–12]. Their activation induces release of cytotoxic molecules such as perforin and granzymes. Activated γδ T cells also secrete cytokines such as interferon-γ (IFN-γ) and tumor necrosis factor-α (TNF-α). These cytotoxic molecules and cytokines induce cancer cell apoptosis. However, γδ T cells comprise only a small percentage of circulating lymphocytes and require stimulation to exert antitumor effects. In this review, we will outline the methods used to stimulate γδ T cells and improve their antitumor effects. We also discuss strategies for clinical application.

#### **2. Phosphoantigens and Nitrogen-Containing Bisphosphonates Stimulate** γδ **T Cells**

Human peripheral blood γδ T cells, which predominantly express the Vδ2 chain paired with the Vγ9 chain, are activated upon recognition of phosphoantigens (PAgs) such as (E)-4-hydroxy-3-methylbut-2-enyl pyrophosphate (HMBPP), which is synthesized in bacteria via isoprenoid biosynthesis [13], and isopentenyl pyrophosphate (IPP), which is produced in eukaryotic cells via the mevalonate pathway [14]. Activation of γδ T cells by PAgs was first reported in the 1990s [15,16]; however, it is unclear how the γδ TCR recognizes PAgs. Butyrophilin subfamily 3 member A1 (BTN3A1) molecules, which are isoforms of the BTN3A (also termed CD277) subfamily, play an indispensable role in activation of γδ T cells by PAgs [17]. BTN3A1, which is expressed ubiquitously on the surface of cells, comprises two immunoglobulin-like extracellular domains and an intracellular B30.2 domain. The precise mechanism by which γδ T cells recognize BTN3A1 is not completely clear, but several studies demonstrate that binding of PAgs directly to a positively-charged pocket in the intracellular B30.2 domain of BTN3A1 recruits the cytoskeletal adaptor protein periplakin and the GTPase RhoB, which increases membrane mobility and induces a conformational change in BTN3A1; the altered conformation is recognized by the γδ TCR [18,19]. Recent studies show that BTN2A1, which binds directly to the TCRs via germline-encoded regions of Vγ9, is also essential to BTN3A-mediated γδ T cell cytotoxicity and BTN2A1 expression at the plasma membrane of cancer cells correlated with γδ T cell cytotoxicity [20,21]. BTN2A1 interacts with BTN3A1, leading to enhance plasma membrane export, and BTN2A1/BTN3A1 interaction is enhanced by PAgs. Anti-BTN2A monoclonal antibodies (mAbs) inhibit BTN2A1 biding to the γδ TCR and modulate γδ T cell killing of cancer cells [21]. These studies demonstrate the potential of butyrophilin subfamily cooperation pathway as a therapeutic target in γδ T cell activation.

In general, the concentrations of PAgs is not high enough to stimulate γδ T cells under physiological conditions; however, tumor cells show upregulated production of PAgs due to metabolic reprogramming, which increases mevalonate pathway activity [22,23]. Moreover, PAgs concentrations can be increased pharmacologically. Nitrogen-containing bisphosphonates (N-bis) such as pamidronate (Pam) and zoledronate acid (ZOL), which are used to treat hypercalcemia or bone metastases of cancer, inhibit the enzyme farnesyl diphosphate (FPP) synthase, which is the rate determining enzyme in the mevalonate pathway [24]. As a result, the concentration of IPP (derived from the upstream FPP synthase metabolite) increases, thereby activating γδ T cells (Figure 1).

**Figure 1.** Mechanism of γδ T cell activation by N-bis. N-bis inhibits FPP synthase in the mevalonate pathway and induces accumulation of IPP. Binding of IPP to the intracellular B30.2 domain of BTN3A1 recruits the cytoskeletal adaptor protein periplakin and the GTPase RhoB, which increases membrane mobility and induces a conformational change in BTN3A1, which is then recognized by the γδ TCR.

γδ T cell-based immunotherapy is classified according to the method used to activate and expand the cells [25]. The first method involves in vivo activation by systemic administration of PAgs or N-bis, along with exogenous interleukin (IL)-2 [26–32] (Table 1). Dieli et al. conducted a phase I clinical trial involving patients with metastatic hormonerefractory prostate cancer. The aim was to examine the antitumor effect of single or combined administration of ZOL and IL-2. Nine patients were enrolled in each arm. Six of the nine patients received combined administration of ZOL and IL-2, but only two of nine patients received single administration of ZOL, and showed a significant long-term shift in peripheral blood γδ T cells toward an activated state in which they produced IFN-γ and perforin; also, the number of activated γδ T cells showed a significant correlation with favorable clinical outcomes [26]. This indicates the importance of the administration of IL-2 to maintain peripheral γδ T cells. Wilhelm et al. reported a pilot study of patients with low-grade non-Hodgkin lymphoma and multiple myeloma; this study involved in vivo activation of γδ T cells by combined administration of Pam and IL-2. The results showed that γδ T cell activation/proliferation and response to treatment were disappointing, with only one of ten patients that received an intravenous infusion of IL-2 on Day 3 through Day 8 achieving stable disease. On the other hand, the next nine patients selected had shown positive in vitro proliferation of γδ T cells in response to Pam/IL-2; when these patients received an intravenous infusion of IL-2 on Day 1 through Day 6, five showed in vivo activation/proliferation of γδ T cells, and three showed a partial response [27]. Therefore, if patients are to have any chance of a clinical responses, they must show positive in vitro proliferation of γδ T cells in response to stimulation with Pam, and IL-2 must be administered immediately after in vivo Pam stimulation. Lang et al. reported a pilot trial of in vivo γδ T cell activation in 12 patients with metastatic renal cell carcinoma (RCC); they used different doses of ZOL in combination with low-dose IL-2. Two patients experienced a prolonged period of stable disease; however, no objective clinical responses were observed [28]. The most common adverse events associated with in vivo-activated γδ T cell immunotherapy are the same as those reported for IL-2 monotherapy; they include fever, fatigue, elevation of liver transaminase, and eosinophilia. These adverse events are usually

grade 1 or 2, meaning that in vivo-activated therapy is well tolerated. However, the clinical benefits appear to be mild to moderate [25]. This problem could be related to anergy and exhaustion of activation-induced γδ T cells. The mechanisms underlying this anergy and exhaustion remain unclear. The second category of γδ T cell immunotherapy involves ex vivo expansion of γδ T cells by PAgs or N-bis, followed by administration of the cultured γδ T cells to the patient (i.e., adoptive immunotherapy) [33–42] (Table 1). The mechanism by which N-bis expands γδ T cells from peripheral blood is as follows: treatment of peripheral blood mononuclear cells with N-bis leads to accumulation of IPP in monocytes because these cells take up N-bis efficiently; the monocytes that accumulate IPP become antigenpresenting cells and stimulate γδ T cells in the peripheral blood [43,44]. Kobayashi et al. conducted a pilot study of adoptive immunotherapy in patients with advanced RCC using autologous γδ T cells stimulated by PAg (namely, 2-methyl-3-butenyl-1-pyrophosphate (2M3B1-PP)). Seven patients were enrolled and all received an intravenous infusion of recombinant human IL-2 plus autologous γδ T cells expanded from their own peripheral blood nuclear cells. All patients had IL-2-related adverse events, which were graded as 1 or 2. The antitumor effects in five patients were evaluated by comparing the tumor-doubling time, assessed by computed tomography (CT), between pre- and post-treatment. Three of the five showed a prolonged tumor-doubling time; however, the other two patients showed a shorter tumor-doubling time. One died within 2 months of γδ T cell administration, and the other showed a shorter tumor-doubling time for liver metastases [33]. In this study, no patient received systemic ZOL. ZOL treatment is important for the antitumor effects of γδ T cells because it inhibits FPP synthase, leading to accumulation of IPP in cancer cells and specific antitumor cytolysis by γδ T cells in a TCR-dependent manner. Kobayashi et al. also conducted a phase I/II study of adoptive γδ T cell immunotherapy in combination with ZOL and IL-2. Enrolled patients had advanced RCC. Eleven patients were enrolled and all received 4 mg ZOL intravenously, followed by administration of autologous γδ T cells starting 2 h after completion of ZOL infusion. Patients then received low-dose recombinant human IL-2 on Day 0 through Day 4. Clinical responses were examined by CT and evaluated using the Response Evaluation Criteria in Solid Tumors. One patient exhibited a complete response, five patients had stable disease (SD), and five had progressive disease (PD) [34]. Nicol et al. reported a clinical study of autologous γδ T cell immunotherapy for various types of metastatic solid tumors (i.e., melanoma, breast cancer, cervical cancer, ovarian cancer, colon cancer, cholangiocarcinoma, and duodenal cancer). Eighteen patients were enrolled. Three of the 14 evaluable patients showed a SD and 11 had PD. Interestingly, this study also examined the migratory pattern of intravenously-infused ex vivo-expanded γδ T cells labeled with radioactive <sup>111</sup>indium oxine (111In) in three patients (two melanoma patients, one colon cancer patient). In all three, labeled γδ T cells migrated rapidly to the lungs and remained there for 4 to 7 h. Cell numbers (estimated by measurement of γ-ray radioactivity in the lungs) decreased slowly, corresponding with gradual migration into the liver and spleen. After 24 h, almost all cells were located in the liver and spleen and virtually no activity remained in the lungs. Moreover, assessment of the number of peripheral blood γδ T cells at multiple time points during the 48 h after γδ T cell infusion showed no substantial change compared with pre-infusion levels. These data indicate that few of the γδ T cells remained in the bloodstream. However, in one melanoma patient of the three patients, the <sup>111</sup>In-labeled γδ T cells appeared to have migrated to the metastatic mass on the left adrenal gland by 1 h after infusion. Maximal activity was seen at the metastatic tumor site at 4 h, and the tracer remained detectable for 48 h [35]. Adoptive immunotherapy using ex vivo-expanded γδ T cells is also safe and well tolerated; however, expanding γδ T cells from some cancer patients is difficult. The reasons for this are unclear. Moreover, favorable clinical outcomes require higher effector (γδ T cells)-to-target cell (cancer cells) ratios (E/T ratio) at the tumor site. Although potent cytotoxic activity against various cancer cells has been confirmed in vitro, there is much room for improvement.


**Table 1.** γδ T cell-based clinical trials.

MM: multiple myeloma; NHL: non-Hodgkin's lymphoma; RCC: renal cell carcinoma; NSCLC: non-small-cell lung cancer; AML: acute myeloid leukemia; Pam: pamidronate; IL-2: interleukin-2; ZOL: zoledronate acid; BrHPP: bromohydrin pyrophosphate; BTN3A: Butyrophilin subfamily 3 member A; mAbs: monoclonal antibodies.

#### **3. Administration of** γδ **T Cells into a Local Cavity Improves the E/T Ratio to Achieve a Maximum Cytotoxic Effect**

The E/T ratio at the tumor site is an important factor that determines cytotoxicity. Administration of effector cells into a local cavity might improve the E/T ratio at the tumor site, making it more likely that γδ T cells make direct contact with cancer cells. Several studies describe administration of γδ T cells into a local cavity, such as the intraperitoneal cavity, enucleated cavity, or intravesical cavity. Wada et al. reported injection of ex vivoexpanded γδ T cells following ZOL administration into the intraperitoneal cavity of seven patients with symptomatic malignant ascites secondary to gastric adenocarcinoma. Two of the seven dropped out of the study after a single injection due to disease progression. In one patient, the bloody ascites became clear and reduced in volume. In another patient, the ascites almost disappeared. The most commonly observed treatment-related adverse events were fever and ZOL-induced hypocalcemia. These events were reversible, and none of the patients experienced abdominal pain or any toxicity related to the intraperitoneal injection of γδ T cells [38]. Nichole et al. reported intracranial infusion of ex vivo-expanded γδ T cells from healthy volunteers into athymic nude mice bearing xenografts of the human glioblastoma (GBM) cell line, U251. Intracranial infusion of γδ T cells led to regression of GBM tumors and improved survival [45]. Intravesical administration of drugs (mitomycin C, adriamycin, or Bacillus Calmette-Guerin) is the standard treatment for bladder cancers. Yuasa et al. implanted a human bladder cancer cell line (UMUC3 cells transfected with the luciferase gene (UMUC3-luc)) into the murine bladder cavity and then administered ex vivo-expanded γδ T cells from healthy volunteers along with 5 µM ZOL by the transurethral and intravesical routes on Day 4 through 8 after cancer cell transplantation [46]. In our previous study, we used an in vivo orthotopic xenograft model to test a protocol based on weekly bladder instillation of γδ T cells, as this is a clinically acceptable schedule [47]. The results of these studies showed that intravesical administration of ex vivo-expanded γδ T cells combined with ZOL inhibits the growth of bladder cancers and prolongs survival significantly. Administration of ex vivo-expanded γδ T cells into a local cavity, rather than systemically, is one strategy that improves the antitumor effects of γδ T cells for clinical application.

#### **4. Other Interactions between** γδ **T Cells and Cancer Cells**

γδ T cells recognize not only PAgs via the γδ TCR, but also stress-associated antigens via the natural killer (NK) group 2 member D (NKG2D) receptor; as for natural killer cells, this method of recognition is MHC unrestricted [48–53]. In 1999, Bauer et al. reported that MHC class I chain-related molecule A (MICA) is a functional ligand that stimulates the NKG2D receptor [49]. In addition to MICA, the MICB and UL16-binding proteins 1–4 (ULBP 1–4) in human NKG2D ligands, as well as interactions between these ligands and the NKG2D receptor, are important for cancer cell recognition and γδ T cell-mediated cytotoxicity [51–53]. Anticancer agents inhibit immune function in cancer patients, mainly through bone marrow suppression [54]. However, recent studies show that some agents amplify the cytotoxic effects of immune cells against cancer cells [55]. Anticancer agents induce the DNA damage response, which in turn upregulates expression of NKG2D ligands [56]. Todaro et al. reported that low concentrations of anticancer agents 5-fluorouracyl and doxorubicin sensitize colon cancer-initiating stem cells to γδ T cell-mediated cytotoxicity via NKG2D receptor:ligand interactions [57]. Lamb et al. showed that temozolomide (TMZ), the main chemotherapeutic agent used to treat GBM, increases expression of NKG2D ligands on TMZ-resistant glioma cells, making them more susceptible to recognition and lysis by γδ T cells [58]. In our previous study, we showed that pretreatment of an orthotopic xenograft model with low-dose gemcitabine upregulates expression of MICA/B in bladder cancer cells and increases the cytotoxic effects of γδ T cells plus ZOL [47]. Molecularly targeted agents also could affect NKG2D ligands. Huang et al. reported that tyrosine kinase inhibitors, sorafenib and sunitinib, markedly increased NK cells cytotoxicity against multidrug-resistant nasopharyngeal carcinoma cells in association with up-regulation of NKG2D ligands, MICA, MICB, and ULBP1-3 [59]. Inhibition of epidermal growth factor receptor (EGFR) pathway also leads to induction of NKG2D ligands. Kim et al. reported that EGFR inhibitors, gefitinib and erlotinib enhanced the susceptibility to NK cell mediated lysis of lung cancer cells by induction of ULBP1 by inhibition of protein kinase C (PKC) pathway [60]. In the γδ T cells field, Story et al. reported that proteasome inhibitor bortezomib significantly increased expression of ULBP 2/5/6 in both acute myeloid leukemia (AML) and T-cell acute lymphoblastic leukemia (T-ALL) cells, and enhanced ex vivo expanded γδ T cell-mediated killing of these cells [61]. Histone deacetylase (HDAC) inhibitors, which are epigenetic agents, are also candidates for combined therapy with γδ T cells. Skov et al. reported that HDAC inhibitors upregulate NKG2D ligands on the surface of several cancer cells [62].

Expression of Fas ligand (FasL) and TNF-related apoptosis-inducing ligand (TRAIL) is upregulated in activated γδ T cells [63]. FasL interacts with CD95, also called Fas or APO-1, which was the first death receptor within the apoptotic chain to be molecularly characterized [64]. CD95 is expressed by various human cancer cells; ligation of CD95 by FasL activates the caspase cascade, which initiates cancer cell apoptosis. TRAIL interacts with five receptors (TRAIL-Rs): death receptor 4 (DR4), DR5, decoy receptor 1 (DcR1), DcR2, and osteoprotegerin [65–69]. Death receptors DR4 and DR5 contain a cytoplasmic region known as the death domain, which enables these receptors to initiate cytotoxic signals when engaged by TRAIL [70]. For these reasons, upregulation of CD95 or death receptors DR4 or DR5 in cancer cells might enhance γδ T cell-mediated cytotoxicity. Several anticancer agents upregulate CD95 or death receptors in cancer cells, thereby sensitizing cancer cells to apoptosis mediated by FasL and TRAIL. Shankar et al. report that paclitaxel, vincristine, vinblastine, camptothecin, etoposide, and doxorubicin upregulate DR4 and DR5 in prostate cancer cells, leading to augmentation of TRAIL-induced apoptosis via caspase activation [71]. Mattarollo et al. reported that etoposide, cisplatin, and doxorubicin upregulate CD95 and DR5 in various cancer cells, and that ex vivo-expanded NK cells kill sensitized targets via FasL- and TRAIL-mediated mechanisms [72]. Indeed, they showed that pretreatment of target cells with anticancer agents increased cytotoxicity to 60–70% (compared with the 5–30% observed when either chemotherapy or NK cells were used alone).

Thus, combination therapy with γδ T cells plus anticancer agents, molecularly targeted agents, and epigenetic agents are a promising strategy to improve the antitumor effects of γδ T cells for clinical application (Figure 2).

**Figure 2.** Interaction between γδ T cells and cancer cells. Anticancer agents, molecularly targeted agents, and epigenetic agents upregulate ligands that activate γδ T cells, thereby increasing cytotoxicity.

#### **5. The Tumor Microenvironment (TME) Limits the Cytotoxicity of** γδ **T Cells by Promoting Their Regulatory Functions, by Secreting Immunosuppressive Cytokines, and by Inhibiting Immune Checkpoint Molecules**

Several studies demonstrate the plasticity of γδ T cells. After activation by PAgs, γδ T cells promote a Th1 immune response by secreting TNF-α and IFN-γ; however, γδ T cells can be polarized into cells with properties similar to those of Th2 cells, Th17 cells, or regulatory T cells (Tregs) [73–76]. Unlike monolayer 2D models and mouse models injected with tumor cells, an actual tumor comprises not only cancer cells but also an extracellular matrix (ECM), stromal cells (such as fibroblasts and mesenchymal stromal cells), vascular networks, and immune cells such as T and B cells, NK cells, and tumorassociated macrophages (TAM). This is the TME. The TME plays a significant role in the subsequent evolution of malignancy [77]. For example, the TME harbors various cytokines, chemokines, growth factors, inflammatory mediators, and matrix remodeling enzymes to facilitate crosstalk between TME-constituting cells [78]; this environment can promote polarization of γδ T cells into Th17-or Treg-like cells that produce IL-17 and transforming growth factor (TGF)-β, which favor cancer cell proliferation [79,80]. IL-17-producing γδ T cells induce angiogenesis and support cancer progression [81,82]. TGF-β secreted by Treg cells can negatively regulate γδ T cells [83]. Moreover, the TME harbors various immunosuppressive cells (Figure 3).

**Figure 3.** Cells in the TME induce polarization and anergy of γδ T cells in various ways (with red font) and some potential strategies to overcome negative effect from the TME are conceivable (in the blue boxes).

Cancer-associated fibroblasts (CAFs), which are recruited to the tumor stroma by growth factors secreted by cancer cells, are key components that maintain an immunosuppressive TME. CAFs produce matrix-crosslinking enzymes and mediate ECM remodeling, resulting in a dense and stiff ECM [84]. The dense and stiff ECM compresses intratumoral blood and lymphatic vessels to increase interstitial tissue pressure, which induces hypoxia and impedes delivery of anticancer agents. The dense and stiff ECM also forms a physical barrier that prevents immune cells from infiltrating the cancer [85]. Provenzano et al. reported that hyaluronic acid (HA) is the primary determinant of the ECM barrier. They showed that enzymatic degradation of HA reduces interstitial tissue pressure to facilitate tumor penetration by gemcitabine, leading to improved antitumor effects in preclinical pancreatic ductal adenocarcinoma transgenic mouse models [86]. HA targeting might permit efficient delivery of γδ T cells to the tumor, thereby improving the E/T ratio on the tumor site. CAFs produce various immunosuppressive cytokines and factors such as IL-6, TGF-β, and prostaglandin E2 (PGE2) [87,88]. IL-6 recruits TAMs and promotes their transition to an immunosuppressive phenotype (i.e., M2 macrophages). CAFs can also inhibit activation of cytotoxic T cells and NK cells directly by expressing inhibitory immune checkpoint molecules such as programmed death-ligand (PD-L)1 and PDL-2 [89].

Myeloid-derived suppressor cells (MDSCs) also play a crucial role in maintaining an immunosuppressive TME. They are converted from immature myeloid cells in the bone marrow by inflammatory mediators released by cancer and immune cells and are recruited to the tumor site through interaction between C-C motif receptors (CCR) and their respective chemokines, such as C-C motif chemokine ligand. They produce different immunosuppressive mediators such as arginase-1 (ARG1), indoleamine 2,3 dioxygenase (IDO), and nitric oxide synthase (iNOS), all of which induce T cell anergy via different pathways [90]. Sacchi et al. reported that MDSCs inhibit IFN-γ production by PAgsactivated γδ T cells and suppress their cytotoxic activity [91]. Several strategies to target MDSCs have been investigated. Blocking migration of MDSCs is one strategy for targeting this cell type. CCR5 plays a key role in migration of MDSCs. The interaction between CCR5 and its ligand CCL5 supports tumor growth and invasion, and migration of MDSCs to the

tumor site; tumor growth and invasiveness are suppressed by targeting the CCR5-CCL5 interaction [92–94]. Inhibiting MDSCs-producing immunosuppressive mediators is another strategy for targeting MDSCs. Serafini et al. reported that sildenafil and tadalafil, both of which are inhibitors of phosphodiesterase-5 (PDE-5), increase antitumor cytotoxic T lymphocyte activity and act synergistically with adoptive vaccine-primed CD8<sup>+</sup> T cell therapy to delay tumor outgrowth in preclinical mouse models by downregulating ARG1 and iNOS activity [95]. Entinostat, a class I HDAC inhibitor, is another candidate agent that neutralizes MDSCs-producing immunosuppressive mediators. Orillion et al. reported that entinostat reduced the expression of ARG1, iNOS, and COX2 by MDSCs, and that the combination of entinostat plus anti-PD-1 antibodies increased survival and delayed tumor growth significantly in several preclinical mouse models [96]. Combination of γδ T cell immunotherapy with PDE-5 inhibitors and HDAC inhibitors is a good strategy for overcoming the immunosuppressive effects of MDSCs.

Tregs, which suppress aberrant immune responses against self-antigens, promote immune evasion of the TME. Infiltration of tumor tissue by a large number of Tregs is often associated with a poor prognosis. They not only exert immunosuppressive activity indirectly by releasing soluble inhibitory molecules such as TGF-β and IL-10, but also directly by inhibiting effector T cells via immune checkpoint receptor cytotoxic T lymphocyte antigen-4 (CTLA-4) and lymphocyte activation gene-3 (LAG-3) [97,98]. Molecules that are relatively specific for Tregs are good candidates for targeting Tregs in combination with γδ T cell immunotherapy. Several studies suggest that an anti-CTLA-4 monoclonal antibody (mAb) predominantly targets Treg cells and strengthens antitumor immune responses [99–101]. Moreover, the clinical efficacy of ipilimumab, a mAb specific for CTLA-4, correlates with a reduction in Treg numbers in tumor tissue [102,103]. CCR4 is expressed predominantly by effector Tregs, which are the most abundant cell type among FOXP3<sup>+</sup> T cells in tumor tissue; in addition, CCR4 ligands produced by cancer cells or by infiltrating macrophages appear to be involved in migration and infiltration of Tregs into various tumor tissues [104,105]. Sugiyama et al. reported that anti-CCR4 mAb treatment selectively depleted effector Tregs and efficiently induced tumor antigen-specific CD4<sup>+</sup> and CD8<sup>+</sup> T cells both in vitro and in vivo [106]. Glucocorticoid-induced TNF receptor-related protein (GITR) is another molecule expressed by Tregs. Ko et al. reported that administration of an agonistic anti-GITR mAb affects tumor-infiltrating Tregs and evokes a potent antitumor immune response, which can eradicate established mouse tumors without eliciting overt autoimmune disease [107].

TAMs also play a pivotal role in the TME by behaving as M2 macrophages; these cells secrete anti-inflammatory factors such as IL-10, TGF-β, and vascular endothelial growth factor (VGEF)-A [108]. These inhibitory cytokines cause cancer cells to become refractory to immunotherapy. Therefore, therapeutic strategies to target TAMs might be effective. Inhibiting differentiation of systemic monocytes once they enter tumor tissue is one strategy to target TAMs. Interaction between CCR2 on monocytes with its ligand (CCL2) induces migration of monocytes from the circulation to the tumor tissue and promotes tumor proliferation. The cytoplasmic protein, FROUNT, binds directly to activated CCR2 and facilitates monocyte infiltration. Inhibition of FROUNT decreased the number of TAMs in an osteosarcoma mouse model [109,110]. Reprogramming of TAMs, i.e., transdifferentiating M2 macrophages to M1 macrophages, is an alternative strategy to target TAMs for cancer immunotherapy. First, M1 macrophages are induced by IFN-γ, and then combined treatment with IL-2 and anti-CD40 induces a switch from an M2 to an M1 phenotype [111]. Moreover, a recent study shows that PD-1 expressed by TAMs inhibits antitumor immunity [112]. Therefore, anti-PD/PD-L1 therapies are expected to have a direct effect on TAMs.

Among these TME-targeting therapies, therapeutic antibodies specific for inhibitory immune checkpoint molecules are an attractive strategy for overcoming the immunosuppressive effects of the TME; this is because various inhibitory immune checkpoint molecules are associated with immunosuppression by various TME-constituting cells. Therapeutic

antibodies specific for PD-1, PD-L1, and CTLA-4, namely immune checkpoint inhibitors, have had a huge impact on cancer immunotherapy over the past decade [113–116]. The combination of adoptive γδ T cells plus immune checkpoint inhibitors is a hopeful strategy for improving their cytotoxicity because PAgs-stimulated γδ T cells express PD-1 [117] and Rossi et al. reported that blockade of PD-1 can boost antitumor effect of γδ T cells against follicular lymphoma [118]. However, we recently reported that PD-1 blockade did not increase the cytotoxicity of γδ T cell against PD-L1 high solid cancer cells and PD-L1 knockdown did not increase the cytotoxicity [119]. The augmentation effect of blockade of PD-1/PD-L1 axis is still controversial. Further studies should investigate how other inhibitory immune checkpoint molecules such as CTLA-4, IDO, and LAG-3, mediate their immunosuppressive effects against γδ T cells, and how these immunosuppressive effects can be circumvented.

#### **6. Cancer Stem Cells (CSCs) Could Mediate Resistance to** γδ **T Cell Immunotherapy**

According to the American Association for Cancer Research (AACR), CSCs are defined as cells within a tumor that possess the capacity to self-renew and to cause the heterogeneous lineages of cancer cells that comprises the tumor [120]. CSCs are a rare cell population within the tumor, but they are spared after conventional therapy because they are resistant and have the capacity to self-renew, ultimately causing tumor relapse and metastasis. Recent studies indicate that CSCs in various solid tumors play an important role in tumor resistance to conventional chemotherapy and radiotherapy [121–123]. Therefore, unsatisfactory clinical responses reported by past clinical trials of γδ T cell immunotherapy against various advanced and recurrent cancers might be due to the presence of CSCs. Moreover, CSCs can modulate immune cell activity by interacting with the TME. Jinushi reported that chemoresistant CSCs promote M2 macrophage differentiation through interferon-regulatory factor-5 (IRF5)- and macrophage-colony stimulating factor (M-CSF)-dependent mechanisms [124]. Schatton et al. reported that malignant melanoma CSCs possess the capacity to inhibit IL-2-dependent T cell activation and support induction of Tregs [125]. In addition, CSCs secrete several immunosuppressive cytokines into the TME, including TGF-β, IL-10, IL-4, and IL-13 [126,127]. CSCs also express high levels of immune checkpoint molecules, which enable them to evade to immune system [128]. Few studies have investigated the relationship between CSCs and γδ T cells. Previously, we generated prostate cancer spheres and used them to examine the cytotoxicity of *ex vivo*-expanded γδ T cells against sphere-derived prostate cancer cells. Sphere-derived prostate cancer cells were resistant to ex vivo-expanded γδ T cells; in addition, their stem cell markers, including CD133, NANOG, SOX2, and OCT4, were upregulating compared with those of parental cells [129]. These results suggest that ex vivo-expanded γδ T cells will not be effective against CSCs. Further research is needed to clarify the mechanisms underlying the resistance of CSCs to human γδ T cells.

#### **7. Novel Forms of** γδ **T Cell Therapy Overcome Current Therapeutic Limitations**

Recently, several strategies have been developed to improve the antitumor effect of γδ T cell immunotherapy. The use of a bispecific antibody, which is typically equipped with a first specificity for an antigen expressed by cancer cells and a second specificity for an activating molecule on effector cells [130], improved the cytotoxicity significantly. Hoh et al. reported that EpCAM/CD3 bispecific antibody enhanced γδ T cell -mediated lysis of hepatoblastoma and paediatric hepatocellular carcinoma cells in spheroid culture models [131]. Oberg et al. reported that ex-vivo expanded γδ T cell administration with the HER2/Vγ9 bispecific antibody significantly reduced the growth of pancreatic cancer and colon cancer in preclinical models [132,133]. They also reported that tribody [(HER2)2xCD16], which comprises two HER2-specific single chain fragment variables fused to a fragment antigen biding directed to the CD16 antigen expressed on γδ T cells and NK cells, enhanced γδ T cells and NK cells-mediated lysis of HER2-expressing tumor cells, such as pancreatic ductal adenocarcinoma, breast cancer, and autologous primary

ovarian tumors [134]. Bispecific antibodies may be promising strategy to overcome current therapeutic limitations. Chimeric antigen receptor-transduced γδ T cells (CAR-γδ T cells) is another novel strategy to overcome current therapeutic limitations. Chimeric antigen receptors (CARs) are usually derived from single-chain variable fragments (scFvs) of antibodies specific for tumor antigens and transduced using viral vectors. Unlike TCRs, which have narrow range of affinities, CARs typically have a much higher and broader range of affinities [135], thus enabling the CAR-γδ T cells to recognize tumor epitopes independently on their TCR. Deniger et al. reported that polyclonal γδ T cells with CD19 specific CAR-γδ T cells enhanced killing of CD19<sup>+</sup> tumor cells compared with CARneg γδ T cells in vitro, and CD19-specific CAR-γδ T cells reduced CD19<sup>+</sup> leukemia xenografts in mice [136]. CAR-T cell immunotherapy has an off-target effect problem. Fisher et al. designed GD2-specific CAR-γδ T cells in order to limit the toxic effects on normal cells. GD2 is abundantly expressed on the surface of neuroblastoma cells and on several other cancer cell types. In this study, γδ T cells recognized the tumor antigen, then the monoclonal antibody against GD2 recognized GD2 and activated the downstream signal domain to exert antitumor effects. Consequently, GD2-expressing neuroblastoma cells which engaged γδ TCR were efficiently lysed, whereas cells that expressed GD2 equivalently bud did not engage γδ TCR were untouched [137]. Currently, several clinical studies have been ongoing (Table 2). CAR-γδ T cells are expected to be a new type of γδ T cell immunotherapy in the future.

**Table 2.** CAR-γδ T cell-based clinical trials.


CAR: chimeric antigen receptor; NKG2DL: natural killer group 2 member D ligand; AML: acute myeloid leukemia.

#### **8. Conclusions**

In this review, we have discussed different ways of activating γδ T cells, along with various strategies aimed at improving their antitumor effects during clinical application. γδ T cell-based immunotherapy is very attractive because these cells show cytotoxic effects against various cancer types, both in vitro and in mouse models. However, clinical trials have reported limited clinical benefit. In vivo activation of γδ T cells by systemic administration of PAgs or N-bis, along with exogenous interleukin (IL)-2, is well tolerated; however, the clinical benefits appear to be mild to moderate, likely due to anergy and exhaustion of activation-induced γδ T cells. However, adoptive immunotherapy using ex vivo-expanded γδ T cells could be achieved by repeated administration of activated γδ T cells, although it is difficult to acquire adequate numbers of activated γδ T cells from some patients. Further research into the mechanisms underlying this problem is needed. Another problem with adoptive immunotherapy conferred by ex vivo-expanded γδ T cells is that systematic intravenous administration of these cells does not achieve a high E/T ratio at the target tumor site. Administration of ex vivo-expanded γδ T cells into a local cavity resolves this problem and is a promising approach to making the most out of their cytotoxic potential. Moreover, pretreatment with anticancer agents, molecularly targeted agents, and epigenetic agents sensitizes cancer cells to γδ T cells by upregulating expression of several stress-induced ligands. Immunosuppression of γδ T cells by the TME and CSCs is less clear-cut, and might operate via multiple mechanisms; however, they affect the immune system via common inhibitory immune checkpoint molecules. Therefore, co-immunotherapy with γδ T cells plus immune checkpoint inhibitors is one strategy that may improve cytotoxicity. Bispecific antibodies and CAR-γδ T cells are novel

strategies which are expected to overcome current therapeutic limitations. Further basic studies of the immunosuppressive effects of the TME and CSCs on γδ T cells, along with clinical studies examining administration into local cavities, combination therapy with anticancer agents, molecularly targeted agents, epigenetic agents, and bispecific antibodies, and CAR-γδ T cell immunotherapy are needed to ensure successful clinical application of γδ T cell-based immunotherapy.

**Author Contributions:** M.M. and T.S. wrote the manuscript and drew the Figures. E.A. and O.U. reviewed the manuscript and finalized it for publication. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by Scholarship donation to the Department of Urology, Kyoto Prefectural University of Medicine.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

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

#### **References**


**Daniel Cacic 1,\* , Oddmund Nordgård 1 , Peter Meyer <sup>1</sup> and Tor Hervig 2,3**


**\*** Correspondence: daniel.limi.cacic@sus.no

**Abstract:** Platelets can modulate cancer through budding of platelet microparticles (PMPs) that can transfer a plethora of bioactive molecules to cancer cells upon internalization. In acute myelogenous leukemia (AML) this can induce chemoresistance, partially through a decrease in cell activity. Here we investigated if the internalization of PMPs protected the monocytic AML cell line, THP-1, from apoptosis by decreasing the initial cellular damage inflicted by treatment with daunorubicin, or via direct modulation of the apoptotic response. We examined whether PMPs could protect against apoptosis after treatment with a selection of inducers, primarily associated with either the intrinsic or the extrinsic apoptotic pathway, and protection was restricted to the agents targeting intrinsic apoptosis. Furthermore, levels of daunorubicin-induced DNA damage, assessed by measuring gH2AX, were reduced in both 2N and 4N cells after PMP co-incubation. Measuring different BCL2 family proteins before and after treatment with daunorubicin revealed that PMPs downregulated the pro-apoptotic PUMA protein. Thus, our findings indicated that PMPs may protect AML cells against apoptosis by reducing DNA damage both dependent and independent of cell cycle phase, and via direct modulation of the intrinsic apoptotic pathway by downregulating PUMA. These findings further support the clinical relevance of platelets and PMPs in AML.

**Keywords:** acute myelogenous leukemia; platelets; microparticles; apoptosis

#### **1. Introduction**

Platelets were originally discovered in the late 19th century as a key player in hemostasis [1]. It is now clear, however, that they serve a broader role in both health and disease [2–7]. Platelets contain many different biologically active molecules, which include proteins [8,9], regulatory microRNAs [10,11], and long RNA sequences, such as ribosomal RNAs and protein-coding transcripts inherited from parental megakaryocytes [12,13]. The long RNA sequences are prone to time-dependent decay [12,14], and correlation with the proteome is weak [13], suggesting only a limited protein synthesis capacity, which may be confined to reticulated platelets [12].

Bioactive substances can be secreted from platelets as paracrine or endocrine factors that are able to modify various cancers [15–17]. These bioactive molecules can also be transferred via platelet microparticles (PMPs), which in turn have been shown to be internalized by many different cancer cells, altering crucial functions of the cells, namely invasiveness, proliferation, and viability [18–20]. The pro-tumoral properties of platelets are further supported by retrospective and observational data showing an association between platelet inhibition and decreased risk for development of cancer, and increased cancer-specific survival [21–24]. However, the mechanism underlying this potential effect remains unknown, and the data from the few prospective studies that have been done are less convincing [25–27].

**Citation:** Cacic, D.; Nordgård, O.; Meyer, P.; Hervig, T. Platelet Microparticles Decrease Daunorubicin-Induced DNA Damage and Modulate Intrinsic Apoptosis in THP-1 Cells. *Int. J. Mol. Sci.* **2021**, *22*, 7264. https://doi.org/10.3390/ ijms22147264

Academic Editor: Angela Stefanachi

Received: 30 May 2021 Accepted: 2 July 2021 Published: 6 July 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 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 (https:// creativecommons.org/licenses/by/ 4.0/).

Acute myelogenous leukemia (AML) is a bone marrow disease affecting hematopoietic stem and progenitor cells [28,29]. The genomic landscape of AML has been thoroughly analyzed and the first study performing whole-genome sequencing in AML was already published in 2008 [30]. AML usually has a lower frequency of somatic mutations than most other cancers [31,32], with a median of 13 different coding mutations per case [33,34]. Despite the low mutational burden, there are large variations in transcriptomic and proteomic signatures in AML cells, compared with healthy hematopoietic progenitors, and even between different subclones [35–37]. Despite increased knowledge in the genomics of AML, treatment strategies have essentially remained unchanged for several decades, with a few exceptions [38]. Curative treatment, which is restricted to younger patients, consists of intensive chemotherapy with consolidating hematopoietic stem cell treatment for high-risk cases. Despite this, median survival is just 11 months when including all age groups [39], underscoring the need for a more profound understanding of progression of the disease and development of treatment resistance, in addition to established genomic mechanisms.

Targeting apoptosis in cancer is a novel treatment strategy that is finally maturing into clinical use. Apoptosis can be divided into two separate pathways, which are interlinked with common feedback mechanisms [40]; the death receptor initiated extrinsic pathway (FAS/CD95, TNFR1, TRAIL-R1, TRAIL-R2, DR3, and DR6), and the intrinsic, or mitochondrial, pathway. Dysregulation of the latter has proven to be an important feature in cancer biology [41]. The regulatory and anti-apoptotic proteins in the BCL2-family are also known to be upregulated in hematological malignancies [42,43]. Hence, numerous drugs that target major apoptotic regulators, such as BCL2 or MCL1, are currently either under development, or have just been approved, to treat a variety of hematological malignancies, including AML [41].

The intrinsic apoptotic pathway is initiated by several factors, including DNA damage or cellular stress, which is accompanied by upregulation of the pro-apoptotic BH3-only proteins (including BAD, BID, NOXA, HRK, BMF, PUMA, BIM), which then activate the effector proteins BAK and BAX directly or through inhibition of anti-apoptotic regulator proteins [44,45]. Upon activation, the predominantly mitochondrial outer membrane (MOM)-bound BAK, and predominantly cytosolic BAX protein, oligomerize in the MOM, leading to cytochrome c leakage from the mitochondria [46,47]. Cytochrome c then forms an apoptosome with apoptotic protease activating factor-1 (APAF1), which recruits procaspase 9, both activating and regulating its function [48,49]. Caspase-9 activates caspase-3, where the intrinsic and extrinsic pathways converge. Caspase-3 has multiple substrates [50], including a caspase-dependent DNase, which leads to DNA degradation upon activation by caspase-3 [51].

Our group has previously shown that co-incubation of the monocytic AML cell line, THP-1, or primary AML samples, with platelet microparticles, protects against daunorubicin (DNR)-induced apoptosis and cell death, at least partially via a decrease in cell activity [52]. We also found that miR-125a and miR-125b levels were elevated in THP-1 cells after PMP co-incubation. These microRNAs have been associated with chemotherapy resistance [53,54]. However, whether the PMP-associated increase in resistance to DNR is caused solely by protection against DNR-induced cell damage, or a modulation of the intrinsic apoptotic pathway regulators, remains unknown. Thus, we sought to further examine the anti-apoptotic effects of PMPs in the monocytic AML cell line THP-1.

#### **2. Results**

#### *2.1. PMPs Offered Protection from Apoptosis Induced by Multiple Agents*

We have previously demonstrated that PMPs increase resistance to DNR-induced apoptosis and cell death [52]. To investigate whether co-incubation with PMPs provided a general anti-apoptotic effect, we compared apoptosis and cell death after treatment with several agents associated with inducing apoptosis, primarily through intrinsic (alantolactone, staurosporine, MG 132), or extrinsic (piceatannol, TRAIL) apoptosis. Co-incubation of PMPs with THP-1 cells decreased the relative frequency of dead and apoptotic cells

**2. Results** 

induced by alantolactone, staurosporine, and MG 132, but not piceatannol (Figure 1). In our experiments 50 ng/mL TRAIL was not sufficient to induce apoptosis in THP-1 cells, but it slightly potentiated the apoptotic effect of piceatannol. Surprisingly, PMP co-incubation increased the relative frequency of dead and apoptotic cells in the case of the combination of piceatannol and TRAIL (*p* = 0.003). However, as this effect was marginal (mean difference of 1.89 percentage points; SD 0.43), it could be biologically insignificant. From these analyses, we suggest that PMPs may provide general protection from apoptosis, but seemingly only against agents that primarily activate the intrinsic apoptotic pathway. of PMPs with THP-1 cells decreased the relative frequency of dead and apoptotic cells induced by alantolactone, staurosporine, and MG 132, but not piceatannol (Figure 1). In our experiments 50 ng/mL TRAIL was not sufficient to induce apoptosis in THP-1 cells, but it slightly potentiated the apoptotic effect of piceatannol. Surprisingly, PMP co-incubation increased the relative frequency of dead and apoptotic cells in the case of the combination of piceatannol and TRAIL (*p* = 0.003). However, as this effect was marginal (mean difference of 1.89 percentage points; SD 0.43), it could be biologically insignificant. From these analyses, we suggest that PMPs may provide general protection from apoptosis, but seemingly only against agents that primarily activate the intrinsic apoptotic pathway.

We have previously demonstrated that PMPs increase resistance to DNR-induced apoptosis and cell death [52]. To investigate whether co-incubation with PMPs provided a general anti-apoptotic effect, we compared apoptosis and cell death after treatment with several agents associated with inducing apoptosis, primarily through intrinsic (alantolactone, staurosporine, MG 132), or extrinsic (piceatannol, TRAIL) apoptosis. Co-incubation

*Int. J. Mol. Sci.* **2021**, *22*, x FOR PEER REVIEW 3 of 17

*2.1. PMPs Offered Protection from Apoptosis Induced by Multiple Agents* 

**Figure 1.** Apoptosis inhibition by platelet microparticles (PMPs). THP-1 cells with or without PMP co-incubation for 24 h were treated with an apoptosis-inducing molecule at a concentration and an incubation time as described in Table S1 (*n* = 4). Relative frequency of dead and apoptotic cells were analyzed by flow cytometry, and gated out in a single gate (annexin V vs. propidium iodide). Data were compared using the paired-sample *t*-test for data pairs. \* *p* < 0.05, \*\* *p* < 0.001. ALA, alantolactone. STS, staurosporine. Pic, piceatannol. TRAIL, tumor necrosis factor-related apoptosis-inducing ligand. **Figure 1.** Apoptosis inhibition by platelet microparticles (PMPs). THP-1 cells with or without PMP co-incubation for 24 h were treated with an apoptosis-inducing molecule at a concentration and an incubation time as described in Table S1 (*n* = 4). Relative frequency of dead and apoptotic cells were analyzed by flow cytometry, and gated out in a single gate (annexin V vs. propidium iodide). Data were compared using the paired-sample *t*-test for data pairs. \* *p* < 0.05, \*\* *p* < 0.001. ALA, alantolactone. STS, staurosporine. Pic, piceatannol. TRAIL, tumor necrosis factor-related apoptosis-inducing ligand.

#### *2.2. PMPs Reduced Both Caspase-8 and Caspase-9 Activation Induced by DNR*

*2.2. PMPs Reduced Both Caspase-8 and Caspase-9 Activation Induced by DNR*  The cytotoxic effect of DNR is commonly associated with an increase in DNA damage, i.e., an intrinsic stimulus. However, it is also suggested to activate the extrinsic apoptotic pathway [55]. To evaluate activation of intrinsic and extrinsic apoptosis after DNRtreatment, we measured levels of active caspase-8 and caspase-9 by flow cytometry, and gated the cells in "lo", "mid", and "hi" populations. In the case of caspase-8, it was not possible to accurately discriminate between the "mid" and "hi" populations, and consequently these populations were gated as one. Our analyses indicated that both caspases were highly activated after DNR-treatment, but this was partially inhibited by PMP co-The cytotoxic effect of DNR is commonly associated with an increase in DNA damage, i.e., an intrinsic stimulus. However, it is also suggested to activate the extrinsic apoptotic pathway [55]. To evaluate activation of intrinsic and extrinsic apoptosis after DNR-treatment, we measured levels of active caspase-8 and caspase-9 by flow cytometry, and gated the cells in "lo", "mid", and "hi" populations. In the case of caspase-8, it was not possible to accurately discriminate between the "mid" and "hi" populations, and consequently these populations were gated as one. Our analyses indicated that both caspases were highly activated after DNR-treatment, but this was partially inhibited by PMP co-incubation (Figure 2A, Figure S3). In addition, fluorescence of the respective caspases were decreased for all subpopulations in PMP co-incubated cells (Figure 2B, Figure S3). The relative decrease in frequency of caspase-8 or caspase-9 "mid/hi" cells associated with PMP co-incubation were equal (Figure 2C; *p* = 0.756). These findings indicated that activation of caspase-8 is important in DNR-induced apoptosis, and is most likely inhibited by PMPs via an upstream mechanism common with caspase-9 activation.

*Int. J. Mol. Sci.* **2021**, *22*, x FOR PEER REVIEW 4 of 17

PMPs via an upstream mechanism common with caspase-9 activation.

incubation (Figure 2A, Figure S3). In addition, fluorescence of the respective caspases were decreased for all subpopulations in PMP co-incubated cells (Figure 2B, Figure S3). The relative decrease in frequency of caspase-8 or caspase-9 "mid/hi" cells associated with PMP co-incubation were equal (Figure 2C; *p* = 0.756). These findings indicated that activation of caspase-8 is important in DNR-induced apoptosis, and is most likely inhibited by

**Figure 2.** Caspase-8 and caspase-9 activation after daunorubicin (DNR)-treatment. (**A**) Active caspase-8 and caspase-9 were analyzed by flow cytometry after 24 h, with or without platelet microparticle (PMP) co-incubation, and an additional 24 h with DNR-treatment at 0.5 µM (*n* = 4). Events were gated as either "lo", "mid", or "hi", according to fluorescence intensity. Data are presented as the relative frequencies of the populations. (**B**) Background corrected mean fluorescence **Figure 2.** Caspase-8 and caspase-9 activation after daunorubicin (DNR)-treatment. (**A**) Active caspase-8 and caspase-9 were analyzed by flow cytometry after 24 h, with or without platelet microparticle (PMP) co-incubation, and an additional 24 h with DNR-treatment at 0.5 µM (*n* = 4). Events were gated as either "lo", "mid", or "hi", according to fluorescence intensity. Data are presented as the relative frequencies of the populations. (**B**) Background corrected mean fluorescence intensity (MFI) of the respective caspases for different populations. (**C**) Ratio of the relative frequencies of the subpopulations "mid" and "hi" combined for respective caspases between THP-1 cells, with versus without PMP co-incubation. Data were compared using the paired-sample *t*-test for data pairs. \* *p* < 0.05. \*\* *p* < 0.001. Casp8, caspase-8. Casp9, caspase-9.

#### *2.3. PMP Co-Incubation Downregulated Pro-Apoptotic PUMA Protein*

To further investigate if PMPs could independently affect intrinsic apoptosis, we analyzed levels of BCL2-family proteins with and without PMP co-incubation, and both with and without DNR. Gating strategy is summarized in Figure 3. For the cell population

only visible with DNR-treatment (P2), levels of BAK, BCL2, MCL1, and PUMA were relatively less increased with PMP co-incubation (Figure 4), when compared to non-DNRtreated THP-1 cells (P1), and the decrease seen in BMF levels was relatively less. We also identified the P1 population in DNR-treated cells, and antibody fluorescence intensity was more or less unaffected, except for BMF, which had a somewhat higher level than the P1 population in non-DNR-treated cells. The relative change in fluorescence intensity accompanying PMP co-incubation was as anticipated, and followed the expected trend of protection from DNR-induced cell damage with PMP co-incubation. For example, the fluorescence intensity of BAK increased with DNR in both groups, but the increase was less with PMP co-incubation than without. However, in the case of PUMA we found a reduced signal intensity with PMP co-incubation in all cell populations, independent of DNR. Thus, we suggest that PMP co-incubation may protect THP-1 cells against DNR-induced cell death, at least partially through downregulation of the pro-apoptotic PUMA protein. visible with DNR-treatment (P2), levels of BAK, BCL2, MCL1, and PUMA were relatively less increased with PMP co-incubation (Figure 4), when compared to non-DNR-treated THP-1 cells (P1), and the decrease seen in BMF levels was relatively less. We also identified the P1 population in DNR-treated cells, and antibody fluorescence intensity was more or less unaffected, except for BMF, which had a somewhat higher level than the P1 population in non-DNR-treated cells. The relative change in fluorescence intensity accompanying PMP co-incubation was as anticipated, and followed the expected trend of protection from DNR-induced cell damage with PMP co-incubation. For example, the fluorescence intensity of BAK increased with DNR in both groups, but the increase was less with PMP co-incubation than without. However, in the case of PUMA we found a reduced signal intensity with PMP co-incubation in all cell populations, independent of DNR. Thus, we suggest that PMP co-incubation may protect THP-1 cells against DNR-induced cell death, at least partially through downregulation of the pro-apoptotic PUMA protein.

To further investigate if PMPs could independently affect intrinsic apoptosis, we analyzed levels of BCL2-family proteins with and without PMP co-incubation, and both with and without DNR. Gating strategy is summarized in Figure 3. For the cell population only

*Int. J. Mol. Sci.* **2021**, *22*, x FOR PEER REVIEW 5 of 17

intensity (MFI) of the respective caspases for different populations. (**C**) Ratio of the relative frequencies of the subpopulations "mid" and "hi" combined for respective caspases between THP-1 cells, with versus without PMP co-incubation. Data were compared using the paired-sample *t*-test for data pairs. \* *p* < 0.05. \*\* *p* < 0.001. Casp8, caspase-8. Casp9, caspase-9.

*2.3. PMP Co-Incubation Downregulated Pro-Apoptotic PUMA Protein* 

**Figure 3.** Gating strategy for intracellular flow cytometry of BCL2-family proteins. Doublets were first discriminated in FSC-A vs. FSC-H plots. P1 represents the population of daunorubicin (DNR)-treated cells gated in FSC-A vs. SSC-A plots that aligned well with non-DNR-treated cells. P2 represents a population generated by DNR-treatment and with increased light scatter. **Figure 3.** Gating strategy for intracellular flow cytometry of BCL2-family proteins. Doublets were first discriminated in FSC-A vs. FSC-H plots. P1 represents the population of daunorubicin (DNR) treated cells gated in FSC-A vs. SSC-A plots that aligned well with non-DNR-treated cells. P2 represents a population generated by DNR-treatment and with increased light scatter.

#### *2.4. Inhibitors of Caspase-9 and BAX Protected Against DNR-Induced Cell Death, but Less so with PMP Co-Incubation*

As PMPs can decrease PUMA protein levels, DNR-induced apoptosis in cells coincubated with PMPs may be less driven by the intrinsic apoptotic pathway. We investigated whether the protective effect of two inhibitors of intrinsic apoptosis, iMAC1 (BAX) and Q-LEHD-Oph (caspase-9), was affected by PMP co-incubation prior to DNR-treatment. We found a lower relative reduction in the relative frequency of dead and apoptotic cells, both for iMAC1 and Q-LEHD-Oph, with PMP co-incubation, which may indicate that caspase-9 activation was a weaker driver in apoptosis (Figure 5A). In addition, inhibiting the activity of caspase-9 or BAX with Q-LEHD-Oph and iMAC1 only yielded a reduction in levels of active caspase-9 in the "NO PMP" setting (Figure 5B). Thus, inhibitors of the

*Int. J. Mol. Sci.* **2021**, *22*, x FOR PEER REVIEW 6 of 17

intrinsic apoptotic pathway were less effective when THP-1 cells were co-incubated with PMPs, suggesting a direct modulation of this pathway.

**Figure 4.** Levels of BCL2-family proteins before and after daunorubicin treatment. (**A**) THP-1 cells were co-incubated with or without platelet microparticles (PMPs) for 24 h before treatment with or without 0.5 µM daunorubicin (DNR) for an additional 24 h. Cells were then analyzed by intracellular flow cytometry. Data were collected as mean fluorescence intensity (MFI) levels corrected for a "no primary antibody" sample of pro-apoptotic BCL2-family proteins (*n* = 5). P1 represents the population determined by viable DNR untreated cells, but also visible in DNR-treated cells, with minimal changes of protein expression. P2 represents the population generated by DNR-treatment. (**B**) MFI levels corrected for a "no primary antibody" sample of anti-apoptotic BCL2-family proteins (*n* = 5). Data were compared using the pairedsample *t*-test for data pairs. \*\* *p* < 0.001. **Figure 4.** Levels of BCL2-family proteins before and after daunorubicin treatment. (**A**) THP-1 cells were co-incubated with or without platelet microparticles (PMPs) for 24 h before treatment with or without 0.5 µM daunorubicin (DNR) for an additional 24 h. Cells were then analyzed by intracellular flow cytometry. Data were collected as mean fluorescence intensity (MFI) levels corrected for a "no primary antibody" sample of pro-apoptotic BCL2-family proteins (*n* = 5). P1 represents the population determined by viable DNR untreated cells, but also visible in DNR-treated cells, with minimal changes of protein expression. P2 represents the population generated by DNR-treatment. (**B**) MFI levels corrected for a "no primary antibody" sample of anti-apoptotic BCL2-family proteins (*n* = 5). Data were compared using the paired-sample *t*-test for data pairs. \*\* *p* < 0.001.

#### *2.*4*. Inhibitors of Caspase-9 and BAX Protected Against DNR-Induced Cell Death, but Less so with PMP Co-Incubation 2.5. PMP Co-Incubation Reduced DNA Damage After DNR-Treatment Independently of Cell Cycle Phase*

As PMPs can decrease PUMA protein levels, DNR-induced apoptosis in cells co-incubated with PMPs may be less driven by the intrinsic apoptotic pathway. We investigated whether the protective effect of two inhibitors of intrinsic apoptosis, iMAC1 (BAX) and Q-LEHD-Oph (caspase-9), was affected by PMP co-incubation prior to DNR-treatment. We found a lower relative reduction in the relative frequency of dead and apoptotic cells, both for iMAC1 and Q-LEHD-Oph, with PMP co-incubation, which may indicate To evaluate the anti-apoptotic effect of PMP co-incubation, we indirectly analyzed double-stranded DNA-breaks (DSBs) through measurement of phosphorylated histone H2AX, or gH2AX, after four h of DNR-treatment. As the process of apoptosis increases DSBs, we first investigated if apoptosis was induced within this time frame. We found that after four h apoptosis was still at the baseline level (Figure 6A). As expected, fluorescence of gH2AX was increased after DNR-treatment in 4N cells compared to 2N cells for both

that caspase-9 activation was a weaker driver in apoptosis (Figure 5A). In addition, inhib-

groups, and the relative frequency of 2N cells was increased with PMP co-incubation (Figure 6B,C). Additionally, the fluorescence of gH2AX was decreased, both for 4N cells, and more surprisingly, for 2N cells with co-incubation of PMPs, compared to the "NO PMP" setting (Figure 6B). These findings indicated that PMP co-incubation protected THP-1 cells against DNR-induced apoptosis by decreasing the amount of DNA damage produced by DNR-treatment, both dependently and independently of cell cycle inhibition. iting the activity of caspase-9 or BAX with Q-LEHD-Oph and iMAC1 only yielded a reduction in levels of active caspase-9 in the "NO PMP" setting (Figure 5B). Thus, inhibitors of the intrinsic apoptotic pathway were less effective when THP-1 cells were co-incubated with PMPs, suggesting a direct modulation of this pathway.

*Int. J. Mol. Sci.* **2021**, *22*, x FOR PEER REVIEW 7 of 17

**Figure 5.** Effects of BAX and caspase-9 inhibitors on apoptosis and caspase-9 activation. (**A**) THP-1 cells were incubated with or without platelet microparticles (PMPs) for 23 h, and then with or without iMAC1 (BAX inhibitor), or Q-LEHD-Oph (caspase-9 inhibitor), for 1 h before treatment with 0.5 µM daunorubicin (DNR). Ratio of relative frequency of dead and apoptotic cells with or without inhibitor was calculated after 24 h *(n* = 3). (**B**) Ratio of relative frequency of caspase-9 mid/hi cells after 24 h of DNR-treatment with or without inhibitor (*n* = 3). Data were compared using the paired-sample *t*test for data pairs. \* *p* < 0.05. **Figure 5.** Effects of BAX and caspase-9 inhibitors on apoptosis and caspase-9 activation. (**A**) THP-1 cells were incubated with or without platelet microparticles (PMPs) for 23 h, and then with or without iMAC1 (BAX inhibitor), or Q-LEHD-Oph (caspase-9 inhibitor), for 1 h before treatment with 0.5 µM daunorubicin (DNR). Ratio of relative frequency of dead and apoptotic cells with or without inhibitor was calculated after 24 h *(n* = 3). (**B**) Ratio of relative frequency of caspase-9mid/hi cells after 24 h of DNR-treatment with or without inhibitor (*n* = 3). Data were compared using the paired-sample *t*-test for data pairs. \* *p* < 0.05.

#### *2.4. PMP Co-Incubation Reduced DNA Damage After DNR-Treatment Independently of Cell*  **3. Discussion**

*Cycle Phase*  To evaluate the anti-apoptotic effect of PMP co-incubation, we indirectly analyzed double-stranded DNA-breaks (DSBs) through measurement of phosphorylated histone H2AX, or gH2AX, after four h of DNR-treatment. As the process of apoptosis increases DSBs, we first investigated if apoptosis was induced within this time frame. We found that after four h apoptosis was still at the baseline level (Figure 6A). As expected, fluorescence of gH2AX was increased after DNR-treatment in 4N cells compared to 2N cells for both groups, and the relative frequency of 2N cells was increased with PMP co-incubation Platelets are now recognized as an important contributor in cancer biology through several mechanisms involving immune evasion, metastasis, and development of cancer microenvironments [15,56–60]. We have previously shown that platelet microparticles increase resistance to DNR in acute myelogenous leukemia cells as a result of decreasing cell activity [52]. Here we provide evidence that this effect is multifactorial. We showed that PMPs protected equally against caspase-8 and caspase-9 activation in DNR-induced apoptosis, and that PMPs decreased DNR-induced DNA damage, not just by inhibiting cell cycle progression. The PMPs also directly modulated intrinsic apoptosis via the downregulation of the pro-apoptotic PUMA protein.

(Figure 6B,C). Additionally, the fluorescence of gH2AX was decreased, both for 4N cells, and more surprisingly, for 2N cells with co-incubation of PMPs, compared to the "NO PMP" setting (Figure 6B). These findings indicated that PMP co-incubation protected THP-1 cells against DNR-induced apoptosis by decreasing the amount of DNA damage produced by DNR-treatment, both dependently and independently of cell cycle inhibition. The anti-apoptotic effect of PMPs was evident with alantolactone, staurosporine, and MG 132, all primarily associated with activation of the intrinsic apoptotic pathway in THP-1 cells [61–63]. On the other hand, PMPs did not protect THP-1 cells against piceatannol or a combination of TRAIL + piceatannol, which are known to activate death receptor 5 and the extrinsic apoptotic pathway [64]. This indicates that PMPs may have broader anti-apoptotic properties, albeit restricted to intrinsic apoptosis. However, it yields no insight into the distinct mechanisms, which could be a common upstream effect on the intrinsic apoptotic pathway, e.g., cell cycle inhibition decreasing induced cellular stress or DNA damage. Interestingly, MG 132 has been shown to induce apoptosis in THP-1 cells arrested in either G1 or G2/M phases, but not when macrophage differentiation is induced [63]. Previously we have shown that PMPs inhibit cell cycle progression, and stimulate differentiation towards macrophages [52]. Thus, the latter could represent an anti-apoptotic mechanism independent of cell cycle inhibition by PMPs.

*Int. J. Mol. Sci.* **2021**, *22*, x FOR PEER REVIEW 8 of 17

**Figure 6.** DNA damage after daunorubicin (DNR) treatment. (**A**) THP-1 cells were co-incubated with or without platelet microparticles (PMPs) for 24 h and treated with 0.5 µM DNR for 4 h before analysis with flow cytometry. Relative frequency of dead and apoptotic cells (*n* = 3). (**B**) Mean fluorescence intensity (MFI) of gH2AX corrected for both an unstained sample and background corrected MFI of representative experiment without DNR (*n* = 4). (**C**) Relative frequency of 2N (G1) and 4N (G2/M) cells (*n* = 4). Data were compared using the paired-sample *t*-test for data pairs. \* *p* < 0.05. **3. Discussion Figure 6.** DNA damage after daunorubicin (DNR) treatment. (**A**) THP-1 cells were co-incubated with or without platelet microparticles (PMPs) for 24 h and treated with 0.5 µM DNR for 4 h before analysis with flow cytometry. Relative frequency of dead and apoptotic cells (*n* = 3). (**B**) Mean fluorescence intensity (MFI) of gH2AX corrected for both an unstained sample and background corrected MFI of representative experiment without DNR (*n* = 4). (**C**) Relative frequency of 2N (G1) and 4N (G2/M) cells (*n* = 4). Data were compared using the paired-sample *t*-test for data pairs. \* *p* < 0.05.

> Platelets are now recognized as an important contributor in cancer biology through several mechanisms involving immune evasion, metastasis, and development of cancer microenvironments [15,56–60]. We have previously shown that platelet microparticles increase resistance to DNR in acute myelogenous leukemia cells as a result of decreasing cell activity [52]. Here we provide evidence that this effect is multifactorial. We showed that PMPs protected equally against caspase-8 and caspase-9 activation in DNR-induced apoptosis, and that PMPs decreased DNR-induced DNA damage, not just by inhibiting Apoptosis induction by DNR is generally believed to be a result of inhibition of topoisomerase II (Top2) enzyme activity, leading to a rise in DNA-Top2 cleavage complexes [65]. The dependence of intact p53 protein for apoptosis induction by doxorubicin, a related Top2 poison, suggests a strong reliance on the activation of the intrinsic apoptotic pathway for this class of chemotherapeutics [66]. However, there is evidence that DNR-treatment upregulates death receptors and activate caspase-8 in multiple leukemic cell lines, thereby

inducing extrinsic apoptosis [55]. In our experiments, caspase-8 and caspase-9 activation was equally inhibited by PMPs, suggesting that PMPs interfere with a common activation mechanism. However, this does not completely rule out a skew in upstream initiation of these pathways, as the levels of active caspase-8 and caspase-9 are also regulated by the downstream caspase-3 and caspase-7 as important feedback mechanisms [40].

We showed that not only was the relative frequency of caspase-9 positive cells lower with PMP co-incubation, but the potency of caspase-9 and BAX inhibitors was also reduced. Both these findings suggest a weaker drive from the intrinsic apoptotic pathway in PMP co-incubated cells, but also correlated with a reduction in the ultimate function of these molecules, which is the inhibition of caspase-9 activation. iMAC1 inhibits conformational activation of BAX, and maybe BAK, without competing with BH3 only proteins [67,68]. The anti-apoptotic effect of iMAC1 is also known to decrease with higher levels of BAX [69], but this should increase chemosensitivity [70], which is the opposite of the effects associated with PMPs. We suggest a common mechanism to explain the reduction in potency of both inhibitors. LEHD (leu-glu-his-asp)-sequence based peptides block the catalytic activity of caspase-9 [71]. iMAC1 will also lead to a decrease in caspase-9 activity by inhibiting mitochondrial outer membrane permeabilization [72]. Thus, both inhibitors ultimately lead to a decrease in the activation of caspase-3, which is not only essential for apoptosis induction, but also for caspase-9 activation in a feedback loop [40]. Thus, if the intrinsic apoptotic pathway is inhibited by PMP co-incubation, the relative contribution of this pathway to caspase-3 activation is reduced compared to the extrinsic pathway, which is also activated by DNR. This should lead to a relative reduction in efficiency of apoptosis inhibition through the intrinsic apoptotic pathway, as extrinsic apoptosis is presumably unaffected by both PMPs and the inhibitors. However, one important caveat for this conclusion is the selectivity of the caspase-inhibitor, which, at least in the older generation inhibitors, is proven to be poor [71]. There are some indications that the second generation inhibitor Q-LEHD-Oph also inhibits caspase-8, but this has not been analyzed in a cell-free system and it was less extensive then the caspase-9 inhibition [73]. Furthermore, our conclusion is supported by results involving two independent inhibitors of separate stages in the intrinsic apoptotic pathway.

The inhibitory effect of PMPs on cell cycle progression is a possible mechanism for increased DNR-resistance, since Top2 poisons are believed to be most effective in proliferative cells [65]. We have previously provided evidence for this, showing that serum starvation of THP-1 cells significantly reduces DNR-induced apoptosis [52]. By measuring gH2AX we showed that PMP co-incubation decreased the level of DNA damage after DNRtreatment. Phosphorylation of histone H2AX is induced by double-stranded DNA-breaks as a DNA damage response [74]. Thus, the level of gH2AX is a widely used proxy for DSBs in biological research [75–77]. As expected, gH2AX levels increased more in dividing 4N cells (G2/M), compared with non-dividing 2N cells (G1) across both groups. As PMPs inhibit cell cycle progression, this would necessarily decrease the level of DNA damage. However, we identified a relative decrease in the signal intensity of gH2AX with PMP co-incubation for both cell phases, suggesting a de facto protective mechanism against the effects of DNR, independent of cell cycle inhibition. Somewhat surprisingly, gH2AX levels were also lower with PMP co-incubation in cells in the G1 cell phase. Previously we have found a decrease in mitochondrial membrane potential associated with PMP co-incubation [52], which may decrease the level of reactive oxygen species (ROS). Significant DNA damage in cells in G1 cell phase is also found in doxorubicin-treated U2OS osteosarcoma cells [78]. This probably has a different etiology compared to the mechanism in the G2/M cell phase and may be explained by an increase in ROS [79,80].

An important question regarding the anti-apoptotic effect of PMPs is if they can directly modulate the apoptotic response. We measured anti-apoptotic (BCL2 and MCL1) and pro-apoptotic (BAK, BMF, and PUMA) BCL2-familiy proteins, both in response to DNR and at baseline, in a "NO DNR" setting. The decreased levels of PUMA associated with PMP co-incubation probably represent a de facto downregulation, as it was present

in all cell populations both with and without DNR. This could be due to increased levels of microRNAs, miR-125a and miR-125b, which are transferred by PMPs [52], and proven to downregulate the protein at the translational level, inducing chemoresistance [53,54]. Furthermore, the "readiness" for activation of intrinsic apoptosis in AML cells has clinical relevance as it is a predictor of outcome with conventional treatment [81]. The other proteins analyzed were also altered, but not in the viable, non-DNR-treated cells, and always in sync with an expected decrease in apoptosis and cell damage associated with PMP co-incubation. Thus, it cannot be stated that these proteins were directly downregulated as a result of PMP-internalization. These differences could be a result of an altered regulation of BCL2-family proteins caused by downregulation of other proteins, such as PUMA. However, they may also stem from a shift in ratio of apoptotic to dead cells, which we did not discriminate. Surprisingly, DNR increased the fluorescence intensity for both the anti-apoptotic proteins tested (BCL2 and MCL1); one would expect a decrease in the level of anti-apoptotic proteins when apoptosis is induced. However, this pattern has been observed for some anti-apoptotic proteins in select leukemic cell lines and is presumably transitory [82].

PUMA is regulated by several factors, including different transcription factors and proteins like forkhead box O (FOXO) and p53 family members [83]. However, these mechanisms may be shared with other pro-apoptotic BCL2-familiy proteins [84], and therefore do not coincide with our observations of isolated PUMA downregulation. PUMA is also post-translationally regulated by phosphorylation and proteosomal degradation, which is proven to be induced by interleukin-3 and HER2 [85,86], but none of these proteins are considered to be a part of the platelet granule or releasate proteome [8,9]. In addition, there are other microRNAs that are present in PMPs, like miR-221 and miR-222 [11], which also are known to downregulate PUMA [87]. However, this has not been investigated in THP-1 or other acute myelogenous leukemia cell lines.

The evidence provided here supplements our previously published work that PMPs have anti-apoptotic properties in acute myelogenous leukemia. This effect could stem partially from inhibition of cell cycle progression and cell activity, making the cells less susceptible to damage induced by chemotherapy. In addition, we showed that PMPs may modulate the intrinsic apoptotic pathway through downregulation of PUMA, as a mechanism independent of cell cycle inhibition. The mechanistic findings from this study were derived solely from one cell line and need to be confirmed in primary AML cells. Nonetheless, translational research with PMPs in AML is warranted, as the indications for platelet inhibition to decrease PMP production, and thus potentially increase chemosensitivity, are further supported.

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

#### *4.1. Cell Line*

The THP-1 cell line was purchased from the American Type Culture Collection (Manassas, VA, USA) and cultured in Iscove's Modified Dulbecco's Medium (IMDM; Thermo Fisher Scientific, Waltham, MA, USA) + 10% FBS (Sigma Aldrich, St. Louis, MO, USA). Culture medium was partially replaced approximately every second day to keep the total cell concentration in the range of 2–6 × 10<sup>5</sup> per mL, and cells were only used in experiments once the exponential growth phase was reached. Cultures were kept for less than three months.

#### *4.2. Platelet Concentrate*

Platelet concentrates pooled from four donors were produced using the automated Tacsi system (Terumo BCT, Lakewood, CO, USA) and were provided by the Department of Immunology and Transfusion Medicine, Stavanger University Hospital (Stavanger, Norway). The platelet concentrations were 0.94–1.06 × 10<sup>9</sup> per mL. Leukocytes were removed by filtration to a residual level of <1.00 × 10<sup>6</sup> . The storage medium for the

platelets was approximately 35% plasma and 65% additive solution (PAS-III; Baxter, Lake Zurich, IL, USA). Written consent was obtained from all donors.

#### *4.3. Platelet Releasate*

Platelet releasate was produced by adding human thrombin (Sigma Aldrich) at a final concentration of 1 U/mL to the platelet concentrates in 50 mL tubes, and incubating for one hour in a 37 ◦C water bath, as described in [52]. The releasates were mixed by gentle shaking every 5 min. To separate the releasate from the clot, the tubes were centrifuged for 10 min at 900 g and the supernatant was transferred to new 50 mL tubes. The samples were stored at −80 ◦C. Fibrin clots that appeared after thawing were removed using a 10 mL serological pipette.

#### *4.4. Platelet Microparticle Production*

Platelet microparticles were isolated as previously described [52]. Briefly, platelet releasate was centrifuged at 15,000 g for 90 min at room temperature, and the supernatant carefully poured off. The PMPs were then resuspended in IMDM + 10% FBS and transferred to cell culture, thoroughly mixing with the cells by pipetting. The final concentration of PMPs was 1.5 × 10<sup>7</sup> per mL culture medium in all experiments. The wells were mixed again by pipetting 2 h after the PMPs were added to the cell cultures.

#### *4.5. Platelet Microparticle Quantitation*

One mL of platelet releasate, washed with 9 mL of Dulbecco's phosphate-buffered saline (Sigma Aldrich), was centrifuged as described above and the supernatant carefully poured off. The platelet microparticles were resuspended in 400 µL of 0.22 µm filtered Annexin V Binding Buffer (Miltenyi Biotec, Bergisch Gladbach, Germany), and 200 µL of the solution was transferred to a second tube. The solution was then stained with 20 µL of Annexin V FITC (Milteny Biotec), and 2 µL of anti-CD61 APC (clone Y2/51; Miltenyi Biotec), or 22 µL of 0.22 µm filtered Annexin V Binding Buffer for a negative control and incubated for 15 min at room temperature. Finally, 278 µL of 0.22 µm filtered Annexin V Binding Buffer and 50 µL CountBright beads (Thermo Fisher Scientific) were added before analysis. Microparticle gates were set using Megamix-PLUS FSC beads (bead size range: 0.3 to 0.9 µm; BioCytex, Marseille, France), according to our previous report [52]. At least 2500 bead events were collected. This, and all other flow cytometric analyses, were performed on a CytoFLEX flow cytometer (Beckman Coulter, Brea, CA, USA) using CytExpert ver. 2.4 acquisition and analysis software (Beckman Coulter). An example of gating strategy for PMP quantitation can be found in Figure S1 (see Supplementary Materials).

#### *4.6. Apoptosis Assay*

Approximately 5 × 10<sup>5</sup> cells per mL THP-1 cells were cultured with or without PMPs for 24 h. The cells were then treated with an apoptosis inductor at a concentration and time interval as indicated in Table S1. Cell viability was analyzed with the Annexin V FITC Kit (Miltenyi Biotec), strictly following the manufacturer's instructions. Dead and apoptotic cells were analyzed using flow cytometry and gated out in a single gate using a dot plot of FITC-A versus PerCP Cy 5.5-A after doublet discrimination. At least 20,000 gated cells were collected. An example of gating strategy can found in Figure S2.

#### *4.7. Apoptosis Inhibition*

For select experiments, after the initial 23 h of incubation with or without PMPs, the THP-1 cells were pretreated with either 20 µM of the caspase-9 inhibitor, Q-LEHD-Oph (Abcam, Cambridge, UK), or 10 µM of the BAX inhibitor, iMAC1 (Sigma Aldrich), and incubated for one hour before adding DNR, as described in the previous section.

#### *4.8. Caspase Activity*

Caspase-8 and caspase-9 activity in THP-1 cells after 24 h of DNR-treatment was measured using the CaspGLOW Fluorescein Active Staining Kit for the respective caspases (Thermo Fisher Scientific). Approximately 5 × 10<sup>5</sup> cells in 0.3 mL IMDM + 10% FBS were incubated with 1 µL of either FITC-IETD-FMK or FITC-LEHD-FMK for 30 min in a CO<sup>2</sup> incubator before washing twice with the supplied wash medium, and analyzing with flow cytometry. Both untreated and treated, but not stained, THP-1 cells were used as controls to determine low, medium, and high caspase populations. At least 20,000 gated cells were collected.

#### *4.9. gH2AX*

Measurement of gH2AX by flow cytometry was performed according to Darzynkiewicz et al. [75]. After PMP co-incubation, cells were fixed with 1% methanol free formaldehyde for 15 min on ice, then fixed and permeabilized in 70% ethanol, and stored overnight. Fixed and permeabilized cells were stained with FITC conjugated antiphospho-Histone H2A.X (Ser139) antibody (clone JBW301; 1 µg/100 µL; Sigma Aldrich), and propidium iodide solution (5 µg/mL; Thermo Fisher Scientific) containing DNase free RNASE A/T1 cocktail (25 U/1000 U per mL; Thermo Fisher Scientific).

#### *4.10. BCL2-Family Proteins*

THP-1 cells were cultured for 24 h with or without PMPs, and an additional 24 h with or without treatment with DNR, before analysis of intracellular proteins using the published protocol by Ludwig et al. [88]. Briefly, cells were fixed and permeabilized using the eBioscience Foxp3/Transcription Factor Staining Buffer Set (Thermo Fisher Scientific). Cells were then incubated with unconjugated antibodies and labeled with the proper conjugated secondary antibodies. A list of the antibodies used, dilutions, and incubation times, can be found in Table S2. A "no primary antibody" sample was used to subtract the background signal. At least 25,000 gated cells were collected.

#### *4.11. Statistical Analyses*

Statistical analyses were performed using the IBM SPSS 26 software (IBM Corp, Armonk, NY, USA). All figures show mean values with 95% confidence intervals. A comparison of means was performed using tests for paired data, or one-sample tests, when appropriate. The data were checked for normality using PP plots, the Shapiro–Wilks test, and the Kolmogorov–Smirnov test. A *p* value < 0.05 was considered significant. "*n*" denotes technical replicates.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/ijms22147264/s1.

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

**Funding:** This work was supported by Western Norway Regional Health Authorities and Bergen Stem Cell Consortium.

**Institutional Review Board Statement:** The study was conducted according to the guidelines of the Declaration of Helsinki, and approved by the regional ethics committee, Regional Etisk Komite Vest (ref 8144) on 16.05.2017.

**Informed Consent Statement:** Informed consent was obtained from all subjects involved in the study.

**Data Availability Statement:** The data presented in this study are available on reasonable request from the corresponding author.

**Acknowledgments:** We would like to thank the Department of Immunology and Transfusion Medicine, Stavanger University Hospital for providing the platelet concentrates used in this study. Graphical abstracted is created with BiroRender.com (accessed on 19 May 2021).

**Conflicts of Interest:** The authors declared no potential conflict of interest.

#### **References**


### *Article* **Cytotoxic Activity against A549 Human Lung Cancer Cells and ADMET Analysis of New Pyrazole Derivatives**

**Agnieszka Czylkowska 1, \* , Małgorzata Szczesio 1 , Anita Raducka 1 , Bartłomiej Rogalewicz 1 , Paweł Kr ˛ecisz 2 , Kamila Czarnecka 2,3 , Paweł Szyma ´nski 2,3 , Monika Pitucha <sup>4</sup> and Tomasz Pawlak 5**


**Abstract:** Two new pyrazole derivatives, namely compound **1** and compound **2,** have been synthesized, and their biological activity has been evaluated. Monocrystals of the obtained compounds were thoroughly investigated using single-crystal X-ray diffraction analysis, FTIR spectroscopy, and NMR spectroscopy. The results gathered from all three techniques are in good agreement, provide complete information about the structures of **1** and **2**, and confirm their high purity. Thermal properties were studied using thermogravimetric analysis; both **1** and **2** are stable at room temperature. In order to better characterize **1** and **2**, some physicochemical and biological properties have been evaluated using ADMET analysis. The cytotoxic activity of both compounds was determined using the MTT assay on the A549 cell line in comparison with etoposide. It was determined that compound **2** was effective in the inhibition of human lung adenocarcinoma cell growth and may be a promising compound for the treatment of lung cancer.

**Keywords:** cytotoxic activity; pyrazole derivatives; MTT assay; ADMET analysis; single-crystal diffraction; FTIR spectroscopy; NMR spectroscopy thermogravimetric analysis

#### **1. Introduction**

There are many different types of compounds, both natural and synthetic, with potential medical properties. Substances, such as imidazoles, oxadiazoles, pyrroles, and many of their derivatives, are well studied and described in the literature [1–5]. The main goal of medicinal chemistry is to synthesize compounds with promising activity and therapeutic agents that exhibit lower toxicity. The search for new, pharmacologically active chemical compounds is related to the modification of existing molecules. A group of compounds that is potentially interesting, due to its structure and biological activity, is those substances containing a pyrazole ring. Pyrazoles exist in many compounds that are used as pharmaceuticals and other active compounds [6–8]. Diseases caused by microbial infection are a serious menace to the health of human beings and often are connected with some other diseases whenever the body system becomes debilitated. The growing incidence of microorganisms that resist antimicrobials is a constant concern for the scientific community. Pyrazoles have always been considered as a scaffold-of-choice in designing novel therapeutic agents because of their anti-inflammatory, anti-tumor,

**Citation:** Czylkowska, A.; Szczesio, M.; Raducka, A.; Rogalewicz, B.; Kr˛ecisz, P.; Czarnecka, K.; Szyma ´nski, P.; Pitucha, M.; Pawlak, T. Cytotoxic Activity against A549 Human Lung Cancer Cells and ADMET Analysis of New Pyrazole Derivatives. *Int. J. Mol. Sci.* **2021**, *22*, 6692. https://doi.org/ 10.3390/ijms22136692

Academic Editor: Angela Stefanachi

Received: 27 May 2021 Accepted: 17 June 2021 Published: 22 June 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 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 (https:// creativecommons.org/licenses/by/ 4.0/).

antihyperglycemic, antipyretic, analgesic, antibacterial, and antifungal properties [9–16]. In this work, we present a synthesis, structural characterization, and investigation of the biological, spectroscopic, and thermal properties of two new pyrazole derivatives. The compounds were also tested for cellular survival through MTT assays on the A549 cell line, which investigated their biological properties.

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

#### *2.1. Synthesis*

The title compounds **1** and **2** have been synthesized in the reaction of 1-cyanophenyl acetic acid hydrazide with isocyanates (butyl and 2,4-dichlorophenyl), according to Scheme 1.

**Scheme 1.** Synthesis route of the obtained pyrazole derivatives.

Compound **1**: 3-amino-*N*<sup>1</sup> ,*N*<sup>2</sup> -dibutyl-5-oxo-4-phenyl-pyrazole-1,2-dicarboxamide. To synthesize compound **1**, 1.75 g of 1-cyanophenylacetic acid hydrazide (0.01 mol) was dissolved while hot and in 15 mL of anhydrous acetonitrile. Then, 1.98 g (0.02 mol) of butyl isocyanate was added to the warm mixture. Stirring was allowed for 7 days at room temperature. The precipitate was then filtered off and dried. The reaction yield was 67%, and the melting point was 131.1 ◦C.

Compound **2**: 3-amino-*N*-(2,4-dichlorophenyl)-5-oxo-4-phenyl-2H-pyrrole-1-carboxamide.

To synthesize compound **2**, 1.75 g of 1-cyanophenylacetic acid hydrazide (0.01 mol) was dissolved in 15 mL of anhydrous acetonitrile. Then, 1.88 g (0.01 mol) of 2,4-dichlorophenyl isocyanate was added. The mixture was kept at room temperature for 5 days. The precipitate was then filtered off and dried. The reaction yield was 81%, and the melting point was 188.9 ◦C.

#### *2.2. Crystal Structure Analysis*

Both structures crystallized in the P21/c space group in a monoclinic crystal system. The crystal data and structure refinement details are summarized in Table 1. Compound **1,** crystallized from CH3OH, has one molecule in the asymmetric *unit* (Figure 1a). One of the butyl fragments shows a disorder. The single crystal of compound **2** crystallized from DMSO. In this case, the molecules of solvents are built into the structure; this involves two DMSO molecules, including one disordered and one water molecule (Figure 1b). Tautomerism was described in an earlier publication from 2014 containing an analogous compound [17]. Compound **2** adopts a keto tautomeric form in a crystalline state. There are typical carbonyl bonds in the structure (C2-O3 = 1.221(4) Å, C9-O10 = 1.242(4) Å). Compound **1** also contains only typical carbonyl groups.


**Table 1.** Crystal and structure refinement data for **1** and **2** compounds.

**Figure 1.** The molecular structure and atom-numbering schemes for: compound **1**—figure (**a**), and compound **2**— figure (**b**), with displacement ellipsoids drawn at the 50% probability level.

The conformation of the molecules is stabilized by intra-molecular hydrogen bonds (S(6)), according to the theory of Bernstein [18]. These are N-H . . . O bonds (N26— H26B· · · O18 and N20—H20· · · O27) (Table 2). The packing of the molecules of the structure **1** is layered (Figure 2). These layers are stabilized by chain hydrogen bonds N26—H26A · · · O25 (C (7)—graph-set) and additionally form dimers N13—H13· · · O27 (R<sup>2</sup> 2 (14)).

**Table 2.** Hydrogen-bond geometry (Å, ◦ ) for **1**.


Symmetry codes: (i) *x*, −*y* + 3/2, *z* − 1/2; (ii) −*x* + 1, −*y* + 1, −*z* + 1.

**Figure 2.** The crystal packing of **1**, viewed along the c axis.

The molecules in structure **2** create gaps into which the disordered solvent molecules are fit (Figure 3). The DMSO molecules form hydrogen bonds of the N-H . . . O(DMSO) type. A water molecule stabilizes the structure with three hydrogen bonds with two **2** molecules and one DMSO molecule (Table 3).

**Figure 3.** The crystal packing of **2**, viewed along the c axis.


**Table 3.** Hydrogen-bond geometry (Å, ◦ ) for **2**.

Symmetry codes: (i) −x + 2, −y + 1, −z + 2 (ii) x + 1, y, z.

#### *2.3. NMR Studies*

It is well known that monocrystals selected for single-crystal X-ray analysis do not always represent the bulk material. Therefore, we additionally performed a validation analysis of the studied materials using the solution NMR technique (Figures S1 and S2 are in the Supplementary Materials). For both compounds **1** and **2,** we recorded series of NMR spectra as <sup>1</sup>H, <sup>13</sup>C, <sup>1</sup>H-1H COSY, <sup>1</sup>H-13C HSQC, and <sup>1</sup>H-13C HMBC, and made full assignments of <sup>1</sup>H as well as <sup>13</sup>C signals. The following NMR parameters were noted for studied compounds: compound **1**, <sup>1</sup>H (DMSO-d6) δ [ppm]: 0.87 (3 × H17, 3 × H24), 1.30 (2 × H16, 2 × H23), 1.44 (2 × H15, 2 × H22), 3.10–3.20 (2 × H14, 2 × H21), 7.20 (H4), 7.28 (H26), 7.37 (H3, H5), 7.49 (H2, H6), 8.10–8.26 (H13, H20) and <sup>13</sup>C (DMSO-d6) δ [ppm]: 13.5–13.6 (C17, C24), 19.2–19.3 (C16–C23), 30.7–31.2 (C15, C22), 39.2–40.1 (C14, C21), 86.0 (C7), 125.6 (C4), 127.3 (C3, C5), 128.2 (C2, C6), 130.2 (C1), 150.5–152.7 (C12, C19), 156.0 (C8), 165.6 (C11); compound **2**, <sup>1</sup>H (DMSO-d6) δ [ppm]: 6.90 (2 × H7), 7.16 (H24), 7.34–7.37 (H23, H25), 7.46 (H15), 7.56 (H22, H26), 7.70 (H13), 8.31 (H16), 10.81 (H5), 11.73 (H1) and <sup>13</sup>C (DMSO-d6) δ [ppm]: 85.9 (C8), 121.6 (C16), 122.9 (C21), 124.9 (C24), 126.6 (C22, C26), 127.1 (C12), 147.1 (C2), 128.0 (C15), 128.2 (C23, C25), 128.8 (C13), 131.4 (C14), 134.3 (C11), 156.3 (C6), 163.2 (C9). Apart from the listed resonances, the residual DMSO, as well as the H2O and HOD signals, were clearly visible on the spectra. The results obtained herein confirmed that both the **1** and **2** samples are homogenous, pure, and stable in DMSO solution. The single-crystal structures presented in the previous section are fully consistent with the NMR results.

#### *2.4. FTIR Spectra*

The FTIR spectra of both compounds confirm their molecular structures. Both spectra (Figures 4 and 5) show several bands of various intensities and shapes that can be ascribed to *ν*(NH) in the range of 3500–3100 cm−1—with the sharpest peaks at 3398, 3328, and 3297 cm−<sup>1</sup> for **1** and 3459, 3308, and 3184 cm−<sup>1</sup> for **2**, respectively. Due to the presence of a butyl group in the **1** molecule, we can observe characteristic bands in the range of 3090–2850 cm−<sup>1</sup> that correspond to alkane *ν*(CH) modes, with peaks at 3081, 3060, 3028, 2954, 2931, and 2869 cm−<sup>1</sup> . In both spectra, we can observe sharp bands in the range of 1730–1680 cm−<sup>1</sup> that result from *ν*(C=O) modes—in the **1** spectrum at 1707 cm−<sup>1</sup> and in the **2** spectrum at 1710 and 1694 cm−<sup>1</sup> . In both spectra, there are several bands present in the 1630–1500 cm−<sup>1</sup> region that can be ascribed to *ν*(CN), *ν*(C=C), and *δ*(NH) modes. Sharp bands in the **1** spectrum at 1461 and 1440 cm−<sup>1</sup> most likely correspond to the *δ*(CH) methyl group modes. When moving to the lower wavenumbers, we can observe bands for both compounds in ranges of 1280–1110 cm−<sup>1</sup> and 790–690 cm−<sup>1</sup> that correspond to *β*(CH) and *γ*(CH) modes, respectively. Several sharp bands that result from *ν*(NN) vibrations can also be observed in the **1** spectrum (1050 and 1016 cm−<sup>1</sup> ) and in the **2** spectrum (1055, 1019 cm−<sup>1</sup> ). The bands that are present in the 870–790 cm−<sup>1</sup> range in the **2** spectrum, but are absent in the **1** spectrum, can most likely be assigned to *ν*(CCl).

**Figure 4.** FTIR spectrum of compound **1**.

**Figure 5.** FTIR spectrum of compound **2**.

#### *2.5. Thermogravimetric Studies in Air*

The thermal decomposition of **1** is shown in Figure 6. This compound is thermally stable up to 125 ◦C. In the first stage of thermolysis, one of the aliphatic chains is destroyed. In the temperature range of 125–175, the exothermic effect on the DTA curve is observed (175 ◦C). When the temperature rises, further destruction of **1** takes place. On the TG curve, there are three mass losses of 8.0% (calc. 7.78%), 19.0% (calc. 19.32%), and 40.0% (calc. 39.68%) within the temperature ranges of 125–175 ◦C, 175–250 ◦C, and 250–525 ◦C, respectively. The final step of decomposition is the burning of organic residues with corresponding exothermic effects on the DTA curve at 680 ◦C. In Figure 7, the TG, DTG, and DTA curves of **2** are shown. Compound **2** starts to decompose at 175 ◦C. The first step of pyrolysis is the destruction of the benzene ring. This process is accompanied by an exothermic peak on the DTA curve at 225 ◦C. The thermolysis of **2** is also a multi-stage and overlapping process. In the temperature range of 175–240 ◦C, the experimental mass loss is 21.0% and it is calculated at 21.23%. The next step is connected with the 52.0% (calc. 52.06%) loss of mass and occurs between 240 and 300 ◦C; on the DTA there are peaks at 270 ◦C, and when the temperature rises above 900 ◦C, the process stops.

**Figure 6.** Thermal decomposition of compound **1**.

**Figure 7.** Thermal decomposition of compound **2**.

#### *2.6. Biological Assays*

In A549 cells, the 50% effective concentration (EC50) for compounds **1** and **2** was found to be 613.22 and 220.20 µM, respectively. The values of the effective concentration after the treatment of the compounds are given in Table 4. It was observed that all synthesized molecules were very active; compound **1** showed much less toxicity than **2**. These results revealed that **2** showed the highest cytotoxicity and the most significant decrease in cell viability relative to the A549 lung cancer cell line. This is very interesting, as both compounds are active in between the activities of the etoposide.

**Table 4.** Cytotoxicity activity at the EC50.


Results are presented as the means ± SD; EC50, 50% inhibition of the cell viability. Statistical significance was assessed using a one-way ANOVA analysis. \* *p* < 0.01 was considered significantly different between cancer and non-cancer cell lines.

#### *2.7. ADMET Analysis*

The pharmacokinetic profile of compound **1** is very promising. ACD/Percepta software indicated optimal human plasma protein binding (74.49%); 4.6 L/kg of distribution volume, which means good distribution to all parts of the human body; and 91.7% of single 50 mg dose bio-availability per os. Compound **2** exhibited more problematic distribution properties: 96.81% human plasma protein binding, 0.34 L/kg of distribution volume, and 36.9% of single 50 mg dose bio-availability per os. Moreover, the prediction results (ACD/Percepta, admetSAR 2.0) indicate the possibility of blood–brain barrier penetration for both structures. The physicochemical profiles of compounds **1** and **2** indicate that both structures are good candidates for drug agents, as they both show the fulfillment of the Lipinski rule [19], the Ghose rule [20], the Egan rule [21], and the Muegge rule [22]. The basic physicochemical properties of compounds **1** and **2** were gathered in Table 5. The analysis results indicate that compounds can affect the pharmacokinetics of other drugs because of their effects on cytochromes P450 isoenzymes. Compound **1** showed inhibition properties for CYP2C19, CYP2C9, CYP3A4, and compound **2** showed inhibition properties for CYP1A2. Both compounds have very promising physicochemical properties for oral bio-availability (Figure 8). Both compounds showed a very low probability of positive AMES test results and hERG inhibition test results. A ProTox II analysis classified both compounds to toxicity class 4 (predicted LD<sup>50</sup> 1000 mg/kg). Moreover, both compounds had very promising results of the detailed prediction of the toxicity profile—compound **1** showed a 0.6 probability of carcinogenicity (1 positive test result out of 17 different predictions), while compound **2** showed a 0.52 probability of carcinogenicity and a 0.64 probability of hepatotoxicity (2 positive test results out of 17 different predictions).

**Table 5.** Basic physicochemical properties of the two compounds.


Log *p* value is an average of 5 prediction algorithms (iLOGP, XLOGP, WLOGP, MLOGP, SILICOS-IT); TPSA—topological polar surface area.

**Figure 8.** Oral bio-availability graph generated using the SwissADME service. The red–coloured zone is physicochemically suitable for oral bio-availability. LIPO—lipophility (−0.7 < XlogP3 < +5.0); SIZE—molecular weight (150 g/mol < MW < 500 g/mol); POLAR—polarity (20 Å<sup>2</sup> < TPSA < 130 Å<sup>2</sup> ); INSOLU—insolubility (0 < logS < 6); INSATU—insaturation (0.25 < fraction Csp3 < 1); FLEX—flexibility (0 < num. of rotatable bonds < 9).

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

#### *3.1. Chemistry*

All of the chemicals used for the synthesis were purchased from Sigma-Aldrich, AlfaAesar, and POCH, and were used without further purification. The FTIR spectra were recorded with an IRTracer-100 Schimadzu Spectrometer (4000–600 cm−<sup>1</sup> ), with an accuracy of recording 1 cm−<sup>1</sup> using KBr pellets. The thermolysis of the compounds in the air atmosphere was studied using TG-DTG-DTA techniques in the temperature range of 25–1000 ◦C at a heating rate of 10 ◦C min−<sup>1</sup> ; TG, DTG, and DTA curves were recorded on a Netzsch TG 209 apparatus under air atmosphere (v = 20 mL × min−<sup>1</sup> ) using ceramic crucibles. Ceramic crucibles were also used as a reference material. All NMR experiments were run at 298 K on a 500 MHz Bruker Avance III spectrometer, which was equipped with <sup>1</sup>H with a <sup>13</sup>C BB probehead (1H-detected experiment) and operating at 500.13 and 125.76 MHz for <sup>1</sup>H and <sup>13</sup>C nuclei, respectively. The samples were prepared in DMSO-d<sup>6</sup> (99.8% + D) from Armar Chemicals. The chemical shifts in <sup>1</sup>H and <sup>13</sup>C were referenced to the methyl groups of DMSO (2.50 and 39.5 ppm, respectively). The <sup>13</sup>C NMR data were assigned by using the standard 2D <sup>1</sup>H-13C NMR correlation techniques, gradientselected heteronuclear single-quantum correlation (gs-HSQC) [23], and gradient-selected heteronuclear multiple-bond correlation (gs-HMBC) [24,25].

#### *3.2. Crystal Structure Determination*

X-ray data were collected at 100 K on an XtaLAB Synergy, Dualflex, Pilatus 300K diffractometer apparatus (Rigaku Corporation, Tokyo, Japan) equipped with a PhotonJet microfocus X-ray tube apparatus (Rigaku Corporation, Tokyo, Japan). Data reduction was performed using CrysAlisPro (Agilent Technologies UK Ltd., Yarnton, UK) [26]. The structure was refined in ShelXL [27]. Molecular plots and packing diagrams were drawn using Mercury [28]. Molecular geometry parameters were computed using PLATON and publCIF [29,30]. The crystallographic information files for the crystal structures are available under the deposition numbers: 2075915 and 2075918.

#### *3.3. ADMET Analysis*

Compound **1** and compound **2** were analyzed using ACDLabs Percepta software version 14.0.0 (Advanced Chemistry Development, Inc., Metropolitan, Toronto, ON, Canada), SwissADME service (Swiss Institute of Bioinformatics, Lausanne, Switzerland, 2021) [31], admetSAR 2.0 service (admetSAR 2019) [32], and ProTOX II service [33] to obtain the computational pharmacokinetic and toxicologic profiles of the tested compounds.

#### *3.4. Biological Assays*

To evaluate the active metabolic cells, the MTT (3-(4,5-dimethylthiazol-2-yl))-2,5 diphenyltetrazoliumbromide) assay was used [34]. In this method, EC<sup>50</sup> (the effective concentration of the tested drug, where a 50% growth reduction is observed in cell growth compared to the untreated control) was used. An MTT assay was performed to test the in vitro cytotoxicity against the A549 cells, which were from a human lung adenocarcinoma that was obtained from the European Collection of Cell Cultures (ECACC, Salisbury, UK). The cells were cultured in Dulbecco's Modified Eagle's Medium (PAN-Biotech, Aidenbach, Germany), 100 units of penicillin/mL (Sigma Aldrich, St. Louis, MO, USA), 100 µg of streptomycin/mL (Sigma Aldrich, St. Louis, MO, USA), 2 mM L-glutamine (Sigma Aldrich, St. Louis, MO, USA), 10% Fetal Bovine Serum (FBS) (Sigma Aldrich, St. Louis, MO, USA), and MTT (3-(4,5-dimethylthiazol-2-yl))-2,5 diphenyltetrazoliumbromide) (Sigma Aldrich, St. Louis, MO, USA). To complete the analyses of the new compounds, the cells were cultured overnight at 37 ◦C with 5% CO<sup>2</sup> in a standard 96-well flat-bottomed plate containing 10<sup>4</sup> cells/well. The following day, the medium was replaced by 100 µL of **1**, **2,** and etoposide added in varying concentrations to the wells. After 24 h of incubation, 50 µL MTT was added to each well for the last 2 h. The final absorbance was measured in analytical wavelengths (570 nm for blue-violet insoluble formazan) using a microplate reader

(Synergy H1, BioTek, Winooski, VT, USA). The viability and cell cycle analysis results were presented as mean ± standard deviation. All assays were performed in triplicate, and the results were obtained in three independent experiments [35].

#### **4. Conclusions**

In summary, etoposide is one of the most commonly used anticancer agents. For many years, it has been the standard therapy for small cell lung cancer, leukemia, lymphoma, germ-cell tumors, and neuroblastoma [36]. Our present findings have shown that both derivatives were very effective in the inhibition of human lung adenocarcinoma cell growth in comparison with etoposide. In conclusion, our present findings showed that the new 3 amino-N-(2,4-dichlorophenyl)-5-oxo-4-phenyl-2,5-dihydro-1H-pyrazole-1-carboxamide **2** (EC<sup>50</sup> = 220.20+/−22.47 µM,) was much more effective in the inhibition of human lung adenocarcinoma cell growth in comparison to compound **1** with 2,4-dichlorophenyl moiety. These results suggest that compound **2** may be a promising molecule for the treatment of lung cancer. In addition, our studies gained new knowledge about pyrazole derivatives.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/ijms22136692/s1. Figure S1. Solution-state 1H NMR (DMSO-d6) of **1** (a) and **2** (b). Figure S2. Solution-state 13C NMR (DMSO-d6) of **1** (a) and **2** (b).

**Author Contributions:** Conceptualization, A.C. and M.S.; methodology, A.C., M.S., K.C., P.S. and M.P.; software, M.S., B.R. and T.P.; formal analysis, A.C. and M.S.; investigation, A.R., B.R., P.K., K.C. and T.P.; data curation, M.S., B.R. and T.P.; writing—original draft preparation, A.C., M.S., A.R. and B.R.; writing—review and editing, A.C., M.S. and B.R.; supervision, A.C., M.S. and P.S.; project administration, A.C. All authors have read and agreed to the published version of the manuscript.

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

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The conducted research was presented for the first time in this publication.

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

#### **References**

