1. Introduction
Poly(3-hydroxybutyrate) (PHB) and polylactic acid (PLA) are two most widely used biodegradable and renewable polyesters, which bear the potential to replace petroleum-derived polymers [
1]. PHB and PLA have also gained vast attention in the investigation of biomaterials-based products [
1,
2,
3].
Poly(3-hydroxybutyrate) (PHB) is a reserve carbon source found in many types of bacteria with chemical structure demonstrated in
Figure 1 [
4,
5]. PHB can be synthesized in various natural environments and it is produced biotechnologically by a series producing strains, such as
Cupriavidus eutrophus [
6] or
Azotobacter chroococcum [
7]. PHB has the ability to degrade into the monomer, 3-hydroxybutyric acid, which is nontoxic and can be removed through metabolism [
8,
9]. PLA is a polyester produced by chemical synthesis from lactic acid. Lactic acid, the monomer of PLA, is produced from sugar or starch using bacterial fermentation or petrochemical methods. PLA can be also hydrolyzed to its monomer
α-hydroxy acid within living organisms. This monomer is then further metabolized through tricarboxylic acid cycle [
10]. Moreover, PLA degradation products are non-toxic and environmentally friendly, making it a first choice for biomedical and industrial applications, such as biomedical implants and food packaging [
11,
12,
13]. However, the PLA degradation products, which are formed during rapid hydrolysis have no time to be taken up by the organism, which causes a drastic pH decrease in proximity to the implant. Chronic tissue irritation caused by reduced pH is considered a serious problem associated with the use of PLA-based polymer implants [
14].
Unlike petro-derived polymers, PHB and PLA exhibit favorable features of biocompatibility, biodegradability, and eco-friendliness, which are very important aspects of biomaterial-based product development. Those features made PLA- and PHB-based materials being exclusively used in many fields ranging from medicine to industrial manufacturing [
15,
16,
17,
18,
19,
20,
21]. Commercially available PLA films and packages are becoming indispensable components for daily life products, while biomedical field has witnessed the great potential of PLA-derived polymers as biocompatible sutures and medical implants. PHB plays a crucial role in designing strategies related to regenerative medicine and tissue engineering due to its superior and desirable characteristics. PHB-based biomaterials have demonstrated their validity for production of conventional medical implantation devices, controlled drug delivery systems, and tissues engineered constructs, including repair patches, scaffolds, adhesion barriers, drug delivery platforms and sutures [
22,
23,
24,
25].
A large number of research has been dedicated to the investigation of these biopolymers’ characteristics, including biodegradability, biocompatibility and mechanical properties, which deepen knowledge of PHB and PLA-based materials [
26,
27,
28,
29]. The ability to promote the use of PHAs will be facilitated by the improved understanding of their biodegradability mechanism [
30,
31]. Despite the presence of a large amount of research that investigates the applications and modifications of PHA, there was a lack of studies regarding the PHA degradation, e.g., surface interaction with cells and dynamic changes of the properties of PHAs. In this research polymer films from PHB and PLA were fabricated for degradation study and cell experiments.
Currently there is a lack of research investigating the impact of culturing cells on polymer films and scaffolds on polymer degradation [
32]. The role of degradation products in cell growth on polymer scaffolds is also poorly understood [
33].
The goal of this project is to study dynamic changes of polymer’s properties, and the interaction between biopolymers and cells during the degradation course.
2. Materials and Methods
2.1. Materials
Bacterial PHB powder was purchased from Biomer (Schwalbach, Germany); PLA and Lipase (isolated from porcine pancreas) were obtained from Merck (former Sigma-Aldrich, Darmstadt, Germany), and bovine serum albumin (BSA) was purchased from Aladdin Industrial Corporation (Shanghai, China).
2.2. Fabrication of Polymer Films
Polymer films were prepared by solution-casting method. Chloroform was used as a solvent for both polymers. A solution of chloroform with a polymer (1.2%) was casted into Petri dishes with a diameter of 9 cm, and Petri dishes were left in the fume cupboard for 24 h until all chloroform evaporated. Finally, films were removed from Petri dishes and cut into small disks (diameter: 1.5 ± 0.2 cm, thickness: 40 ± 5 μm, weight: 9 ± 3 mg). A polymer mixture of PLA and PHB (50:50) was made by mixing solutions of PHB and PLA with thorough mixing.
2.3. Enzymatic and Non-Enzymatic Hydrolytic Degradation
Non-enzymatic degradation was conducted in phosphate buffer saline (PBS) Merck (former Sigma-Aldrich, Darmstadt, Germany) at pH of 7.4. For enzymatic degradation, lipase was added to PBS with a concentration of 0.25 mg/mL. All solutions were stored in a shaker-incubator (37 °C, 150 rpm). To maintain the buffer quality, free PBS buffer and lipase-containing PBS buffer were changed every 3 days. Films were examined at 4 different degradation points (1 week, 2 weeks, 4 weeks and 6 weeks). To prevent bacterial contribution to the degradation of polymers, sodium azide (2 g/L) was added to the buffer solution.
2.4. T3 Cell Seeding Experiments
Murine NIH 3T3 fibroblasts (Biolot, Russia) were used to study the interaction between polymer films and cells at different degradation stages. Cell growth medium was made according to the protocol: Dulbecco’s modified eagle medium (DMEM, PanEco, Moscow, Russia) supplemented with 10% fetal bovine serum (FCS, Biological Industries, Beit-Haemek, Israel), 100 U/mL penicillin (PanEco, Moscow, Russia). The experiment consisted of two parts. For the first part, the experiment was carried out on cell culture plate (24 wells with a diameter of 1.5 cm). Fresh-made films were immersed in ethanol for sterilization (for at least 1 h), followed by washing with sterile PBS. Then, films were flattened and fixed to the bottom of wells. For cell seeding, we added 200 μL of cell suspension (about 2 × 105 cells) and 800 μL of cell growth medium into each well. Finally, the cell culture plate was stored in the CO2 incubator (Sony, Minato, Tokyo, Japan) at 37 °C. Degradation of films and the growth of 3T3 cells were observed at 2 different time points: 1 week, 2 weeks. The second cell experiment was performed on a 96-well plate, and the samples included fresh-made films, films after 2-week’s and 6-week-’s degradation. After sterilization, each film was cut into 3 identical pieces, followed by fixation into 96-well plate. For each well, 100 μL of 3T3 cell solution (2 × 105 cells) together with 200 μL cell growth medium was added for cell seeding. The plate was incubated at 37 °C with CO2 rate 5.0%, and the cell viability test was conducted at 3 different time points (1d, 3d and 5d). The approvals for all experimental procedures and ethical guidelines were issued by the ISO 10993-1:2009.
2.5. Weight Loss Analysis of Polymer Films
Weight loss measurement were carried out at 4 different degradation points (1 week, 2 weeks, 4 weeks and 6 weeks). The changes in the weight of the films during the degradation were determined gravimetrically on the AL-64 scales (Max = 60 g, d = 0.1 mg, Acculab (Sartorius Group), Göttingen, Germany). At each time point, films were taken out of the degradation buffer and washed with distilled water, followed by immersing in 0.1% SDS solution for 24 h. After that, films were washed with distilled water and then dried out. Finally, the weight of each film was measured using microbalance. The weight loss was calculated using the following equation:
where W
0 is the initial sample weight and W
d represents measured sample weight after degradation.
2.6. Particle Distribution Analysis of Film Degradation Products
Samples of PHB, PLA, and PHB-PLA polymer films of the same weight were washed twice and incubated in milli-Q water at 37 °C with constant stirring. After 7 and 14 days of incubation, the films were removed, and the remaining solutions were centrifuged twice for 5 min at 10,000 rpm. The resulting supernatants were analyzed for polymer degradation products by Dynamic light scattering using a Zetasizer Nano ZS (Malvern Panalytical, Malvern, UK). The distribution of nano- and microparticles size was recorded using Zetasizer Software in four independent replicates.
2.7. Molecular Weight (Mw) Analysis
Ubbelohde viscometer was used to measure the viscosity of PHB, PLA and PHB/PLA blend. Polymer film was dissolved in 7 mL chloroform, and the solution was transferred into viscometer. The experiment was carried out in water bath at 30 °C (
Supplementary, Figures S1 and S2). For each sample, the viscosity measurement was repeated 3 times. Molecular weights of PHB and PLA were calculated using Mark-Houwink-Kuhn-Sakurada (MHKS) equation [
7]. The molecular weight of PHB/PLA blend was approximated using the known ratios of PHB and PLA and the rate of their decomposition (
Supplemental Materials, Figures S1 and S2).
2.8. Differential Scanning Calorimetry (DSC)
Thermal properties of PHB, PLA and the blend were measured via differential scanning calorimetry using a DSC 204 F1 Phoenix (Netzsch, Selb, Germany) equipment. The samples were heated from 25 to 200 °C at a heating rate of 10 °C/min in nitrogen environment. The onset and peak temperature of the change in heat capacity was designated as the
(onset) and
(peak). The accuracy of obtained values did not exceed 1 °C for the temperature measure and 2 J/g for the melting enthalpy. The crystallinity of biopolymer component (Xc) was calculated by the following equation [
34]:
where
(PHB) and
m (PLA)represent the theoretical values for the thermodynamic melting enthalpy of completely crystallized PHB (146.6 J/g) and PLA (93.1 J/g), respectively. ΔHm is the melting enthalpy that was calculated from DSC curve. (PHB) and (PLA) are the weight fractions of PHB and PLA in the blend. All calculations were performed for the second heating cycle. Data was presented as the average of three measurements,
p values
< 0.05 were considered statistically significant.
2.9. Scanning Electron Microscopy
To investigate the changes of surface structure and cell distribution, scanning electron microscopy (SEM) and fluorescence microscopy were used. For the SEM’s sample preparation standard dehydration process was conducted using different concentrations of ethanol (30%, 50%, 70% and 96%). After that, Hexamethyldisilazane (HMDS) was used to replace ethanol during sample drying. Finally, the specimens were mounted on metal stubs; then stubs were coated with silver in a sputtering device for 15 min at 15 mA (IB-3, Giko, Fukuoka, Japan) and examined under a scanning electron microscope JSM-6380LA (Jeol, Akishima, Tokyo, Japan). For fluorescence microscope films were cut into small pieces, followed by their immersion in PBS buffer containing Calcein for 15 min. After that, samples were removed from solution and washed by PBS. Finally, the samples were analyzed using fluorescence microscope (blue filter) ZEISS Axio Lab A1 (Zeiss, Oberkochen, Germany).
2.10. Cell Proliferation Assay XTT Test
In order to determine the viability of NIH 3T3 fibroblasts seeded on films XTT cell proliferation test was used. Films were removed from the medium and cut into identical pieces. The film fragments were then transferred into a 96-well plate. For each well 100 μL of DMEM and 50 μL of reaction solution of Cell Proliferation Kit XTT (Biological Industries, Israel: 2% activation solution plus 98% XTT reagent solution was added followed by the plate incubation for 2 h at 37 °C. Film fragments were taken out of the wells after 2-h incubation. After that, the measurement of the absorbance of the samples was carried out using Zenyth 3100 Microplate Multimode Detector (Anthos Labtec Instruments GmbH, Wals, Austria) at a wavelength of 450 nm; 630 nm wavelength was used as a reference absorbance. Then the calibration plot “cell number—absorbance at 450 nm” was built using the known number of the same cells counted by light microscope with the micrometer scale [
35].
2.11. Statistical Analysis
Each experiment was conducted from 2 to 3 times with 3 samples for each polymer material (PHB, PLA, PHB/PLA). The non-parametric Kruskal–Wallis test was employed for the statistical evaluation of data using the software package SPSS/PC+ Statistics™ 12.1 (SPSS: An IBM Company, Armonk, NY, USA). The obtained data were represented as mean ± SD (standard error of the mean), and was considered significant for p < 0.05.
4. Discussion
Weight loss is one of the most conspicuous features observed in polymer degradation, indicating the polymer destruction.
Figure 2 reveals that PLA has a greater weight loss than PHB, which is in accordance with literature. It is connected with that PHB has high crystallinity [
28,
37]. Furthermore, weight loss of PHB has no significant differences between its enzymatic and non-enzymatic degradation.
Molecular weight loss serves as important indication of polymer degradation.
Figure 3 reveals that enzymatic degradation induced a greater molecular weight loss than non-enzymatic degradation, which can be explained by involvement of lipase in accelerating the degradation rate. Researches also confirmed that enzymes, like lipase and depolymerase, are more efficient in contributing to the breakdown of polymer chains [
38,
39]. Although PLA had a smaller initial molecular weight value than PHB, it exhibited a higher loss value (54%) after 6 weeks of enzymatic degradation, where according to the literature, PLA is more prone to degradation than PHB [
26,
28].
The kinetic model parameter (k
D) demonstrated interesting data. During PHB and PLA homopolymer decomposition in a lipase solution, the kinetics curves had two distinct stages of molecular weight change. At the first stage (the first 2 weeks of degradation) the rate of degradation was twice as high as during the following 4 weeks. This can be explained by the presence of an amorphous component in semicrystalline polymers. It was shown earlier that the amorphous component decomposes much faster than the crystalline one [
40,
41]. Autocatalysis is observed during the decomposition of PLA in a phosphate buffer (
Table 2). The absence of this phenomenon during the enzymatic decomposition of PLA is explained by the higher rate of decomposition, at which autocatalysis is not observable. The static state of PHB between 4 and 6 week’s degradation might result from the impediment imposed by the crystallized inner part, which was exposed to the buffer after depletion of amorphous outer regions. It was suggested that further crystallinity analysis may help to explain the unchanged state.
Thermal properties, such as crystallinity, serve as an important indicator of polymer degradation, where the properties changes vary among different polymers and copolymers [
28]. It was reported that the rate of erosion of melt-crystallized films significantly decreased as the degree of crystallinity increased [
39]. DSC analysis revealed (
Table 2) that PHB remained at a constant melting temperature both in blend and pure PHB, which was also in accordance with literature [
42]. The crystallinity of pure PHB displayed relatively small change after 6 weeks’ degradation, suggesting that PHB films still maintained a relatively ordered crystalline structure, which further suggests that PHB had the lowest weight loss and degradation rate compared with the blend and PLA (
Figure 7). Pure PLA exhibited reduced crystallinity after 6 weeks’ degradation, which may indicate the erosion of crystalline phase.
Previous research also confirms that enzymes came into effect when the ordered structures of PHB were dismantled by initial degradation [
43,
44]. The slow degradation of PHB indicates that high polymer crystallinity restricts the effectiveness of enzymes, which slows down the degradation process. Enzyme cannot penetrate highly ordered structures of PHB films at this stage but can penetrate the amorphous phase of PHB and hydrolyze polymer chains. The high molecular weight loss of PHB is associated with cleavage of PHB chains in amorphous phase, which was earlier demonstrated by the work of Zhuikov et al. [
45]. However, this effect of less susceptibility to degradation by enzymes at a later stage is also present for PLA films, whereas the rate of its degradation is much higher (
Figure 6b).
The blend fabricated through mixing PHB and PLA was expected to demonstrate weight loss that would be an average of the value of its primary components PHB and PLA. However, the experimental results showed that blend underwent the highest weight loss after 6 weeks’ enzymatic degradation. It was observed that the PHB/PLA films had a pronounced roughness and porosity after 6 weeks’ enzymatic degradation, which supports the fact that the blend films degraded faster than PLA films following 6 weeks of enzymatic degradation. The unexpected data were obtained upon analysis of the change in the PHB/PLA blend molecular weight. The blending of PHB with PLA reduced the degradation rate and, turning out to have the smallest rate under both enzymatic and hydrolytic degradation. Unlike k
D of PHB and PLA homopolymers, the k
D of PHB/PLA blends stayed the same during both during enzymatic degradation and during lipase-free hydrolysis (
Table 1). The curve had one relatively straight section (
Figure 6c). This behavior can be attributed to a possible interaction between PHB and PLA within the mixture. The degradation rate of the blend is about 4-times less than the rate of the individual homopolymers at the first stage. Notably, the crystallinity degree of the polymers in the blend was also observed to be lower (
Figure 2). The blend’s k
D value is close to the PLA k
D value at the second stage of its degradation, which suggests that PLA could contribute to the blend’s degradation at least during the late stages of the course of the experiment (3–6 weeks). The analysis of blend degradation curves showed that the correlation coefficients of both nonautocatalic and autocatalic models were quite close (
Table 2), which suggests that the contribution of autocatalysis cannot be excluded. Thus, regarding the curve of PHB/PLA blend weight loss during its enzymatic degradation (
Figure 1), we can assume that this effect may be due to the removal of low-molecular degradation products, whereas they are retained in the polymer matrices of PHB and PLA. This can lead to a more uniform and a less pronounced degradation process in the blend.
PHB/PLA blend possessed a more complicated pattern with an increase in crystallinity in PHB component, which may have signified a temporal positive impact of the degradation stages of the initial erosion of amorphous phases on the crystallinity. According to the DSC curve (
Figure 5), the PHB and PLA polymers were immiscible. The characteristic peaks are well distinguishable for each of the polymers. The glass transition temperature and the melting temperature for PLA were found to be 48.8 °C and 146.6 °C, respectively. Moreover, the peak of PLA crystallization (95.1 °C) was observed. This peak is not always visually accessible upon use of polymer mixtures [
26]. The typical melting point peak of PHB was at 175.4 °C, and in contrast to the PLA, the crystallization temperature via cooling was at 70.9 degrees. It was shown also that PLA and PHB are immiscible. The blending of these polymers may compromise the ordered solid structures of polymers, making the blend vulnerable to the enzymatic attack [
26,
46].
These data indicate that the film represents inclusions from PHB and PLA, located randomly throughout the volume. The PLA component in blend had lower crystallinity in enzymatic degradation than that in non-enzymatic degradation, which accounts for highest weight loss of the blend upon enzymatic degradation. It is worth noting that in comparison to pure polymers, both PLA and PHB components in blend experienced a decrease in crystallinity, which is related to the immiscibility and structure disruption between PLA and PHB. In this research, we speculated that following initial erosion of ordered regions, enzymes were capable of penetrating inside the regions with greater fragility and immiscibility that were formed by combination of PHB and PLA, therefore accelerating the degradation rate. Upon suggesting that the polymers are in fact immiscible, this implies a potential increase in the polymer molecules’ surface area, which furthermore may elevate the degradation rate. Surprisingly, this immiscible nature of PHB/PLA blend leads to enzymatic degradation with uniform rate, since enzymes better penetrate into the unorganized polymer blend matrix, especially at the late stage of degradation process, in comparison with the pure polymers, their components: PHB and PLA.
Analysis of the particulate degradation products from polymer films after first week showed very low concentrations of the released particles. The highest concentration was achieved only during PHB films degradation, which was in agreement with our results. This explains the largest decrease in PHB molecular weight in the first week of degradation. After the second week of incubation in water, it became possible to detect nanoparticles. The highest concentrations correspond to particles with a size of 340 nm for PHB, 300 nm for PLA, and 340–530 nm for a composite. The particles of about 200 nm were discovered after the first week of PHB incubation. During this time polymers of this type not have enough time to undergo the decomposition process due to their hydrophobicity. Therefore, the most accurate explanation for this observation is the washing out of oligomers and unbound polymer from the bulk of the product. During the two weeks of incubation, which in turn elevates the particle fraction and leads to the formation of the particle peaks of the order of 10 nm. Particles with the size over 5500 nm are considered an artifact, arising from an insufficient centrifugation, as well as the initial fraction of the polymer, which gives rise to smaller nanoparticles.
Films seeded with cells may have different changes in properties compared to pure films. The molecular weight analysis showed that the growth of 3T3 cells on PLA and blend helped to accelerate the degradation rate, which may have been influenced by the cells’ metabolism. Moreover, 3T3 cells had a wider distribution across PLA’s and the blend’s surface (confirmed by images obtained from SEM and fluorescence microscope). The PHB degradation has shown an opposite trend, where no significant difference in cell experiment and degradation experiment, which was confirmed by a relatively unaffected PHB surface. It was suggested that the highly ordered structure of PHB restricts the possible effects imposed by cell metabolism in the initial stage of degradation. Besides, it can also depend on the overall number of cells grown on films at each time point, which might require further investigation to reveal the distinctive pattern of PHB.
It can be observed that PLA had greater molecular weight loss in cell experiment than in non-enzymatic degradation, while molecular weight loss of PHB and PHB/PLA blend in non-enzymatic degradation was higher than in cell experiment (
Table 4).
The molecular weight loss of enzymatic experiment was higher than cell experiment. A possible explanation for this effect is that upon enzymatic decomposition, the enzyme is provided with a large polymer area, which leads to an accelerated degradation. Cells also accelerate the degradation of polymers but act locally. In contrast to cell-based components, a higher PHB degradation rate in the non-cell (PBS) can be explained by the facilitated leakage of the solutions into the polymer matrix. Cells spread out on the surface of the polymer and, and show a smaller effect on it due to the hydrophobicity of the polymer. In cell investigation PHB/PLA blend also demonstrated more uniform and slower degradation process than PHB and PLA that can be explained by the uneven ultrastructure of the polymer blend matrix and, therefore, its better availability for cleavage by cellular enzymes.
Cell viability test is widely used for evaluation of cell proliferation on polymer films [
35,
47]. Higher cell viability value indicates better cell proliferation and adhesion. The blend and PLA had higher cell viability values after 2 weeks incubation, which implies better cell adhesion and proliferation (
Figure 9). It is also suggested that PLA and blend films display better interaction with 3T3 cells, which is confirmed by images obtained from SEM. This difference in cell viability between polymers may be related with a combination of favorable properties of polymer film surface, such as hydrophilicity and microstructure, which requires further investigation.
Cell experiment on degraded films revealed that there is no significant difference in cell viability between 1d and 3d’s incubation, which may be explained by a small size of films and insufficient incubation time. After 5d’s incubation, it was observed that films with different durations of degradation exhibited distinctive cell viability values. For PHB, fresh films and films after 2 weeks’ degradation had higher cell viability than films with 6 weeks’ degradation, indicating that the degradation duration may have shown a negative correlation with cell proliferation. PLA had a similar cell viability pattern as PHB after 5d’s incubation the only exception with the films undergoing a 6 week enzymatic degradation, which showed higher cell viability. It was suggested that there is no linear correlation between degradation duration and cell viability. Interestingly, the blend displayed relatively stable cell viability in spite of films with different degree of degradation, which indicates that the biocompatability of PHB/PLA blend was retained after 6 week degradation. This effect is also correspondent with more uniform degradation process of the blend.
It was reported in literature that the surface structure of polymers can influence proliferation and adhesion of cells and also that it it depends on specific microstructures of certain biopolymers [
18,
47]. Images obtained from scanning electron microscope revealed that the blend had higher degree of surface erosion (
Figure 11c), which can be explained by the fact that the polymer blending disrupted the ordered structure of pure PLA and PHB, making the blend prone to degradation. PHB retained its relatively smooth and intact surface, which can be attributed to its high crystallinity and high viscosity. The difference in cell distribution may be related to the polymer’s initial surface properties, which influenced cell distribution pattern during the experiment. Meanwhile, it also could be a result of interaction between cells and polymers, which impose a mutual effect on both degradation course and cell proliferation.
The micro- and nanoparticles’ analyses during the degradation of polymers in milli-Q water showed presence of polymer particles in solution with the following size distributions: PHB: 3, 10, 340 nm. PLA: 0.9, 300 nm and for PHB\PLA: 4.1, 340–530 nm. Particle sizes of 300–530 nm can correspond to polymer particles that are not bound to the polymer matrix and released from the bulk of the polymer film. Nanoparticles with size range of 0.9 to 4.1 nm were also observed and were suggested to be a decomposed part of the polymer.
According to the results (
Figure S5), cell proliferation on centrifuged samples did not differ from cells in the control medium, which suggests that the products of PHB degradation are not toxic for them, whereas non-centrifuged PHB powder dispersion in medium inhibited cell proliferation. However some authors implied that PHB nanoparticles do have a toxic effect [
33].
The fluorescence microscopy data shows that cells exhibited better proliferation on fresh made films after 5d’s incubation, which indicates that all polymer films were biocompatible. It was also observed that degraded films of blend and PHB had different cell distribution patterns compared to PLA, which were characterized by a combination of large and small colonies. We speculated that this difference may be related to surface structure and molecular weight. The blend suffered severe surface erosion after 6 weeks’ degradation, which disrupted and restricted cell proliferation to isolated areas with different scales. Although PHB did not experience a high degree of surface erosion after 6 weeks’ degradation, it experienced the highest molecular weight loss, which could be a factor causing the scattered pattern of cell distribution.
Surprisingly, the uneven distribution of cells on polymer films can also be the reason for their slower and more uniform degradation. It was suggested also that the information about cell distribution patterns can be of great benefits for designing 3D-scaffolds seeded with cells for tissue and organ repair.
In bacterial cells, PHB is accumulated in special granules—carbonosomes. It was shown that PHB in such granules is at amorphous condition, which is maintained by a series of PHB-binding proteins [
48]. Such an amorphous conformation of PHB allows the bacterial cell to use this biopolymer as a carbon source for its vital functions. The blending of PHB with PLA can be, with some reservations, a prospective tool to mimic a natural irregular conformation of PHB in bacterial carbonosomes. This approach can help to achieve the desired kinetics of polymer biodegradation for its application in regenerative medicine and tissue engineering.