1.0 Introduction
In recent years, new APIs have increasingly reduced in solubility and is estimated that approximately 70% of new chemical products possess low solubility in both aqueous and organic environments.1 This has led to one of the main confronts of the pharmaceutical industry, proving that effective drug delivery systems are as essential as pharmacodynamics and efficacy of a drug.
The bioavailability of a drug, is the measurement of how much administered drug reaches systemic circulation. The wide variety of physicochemical factors which affect bioavailability are therefore considered in pre-/post formulation to achieve efficient drug delivery. Ideally, for oral preparations, high cell permeability and high solubility, effectively attains a desired high bioavailability.
Figure 1.1 FDA approved schematic of the Biopharmacetuical Classification System2
The biopharmaceutical classification system (BCS) was approved by the FDA based on the previous research of Amidon et al.3 The system is a schematic of barriers limiting bioavailability, where Class 1 has the highest bioavailability, Class 4 the least.
1.1 General issues with drug delivery
In aqueous environments, hydrophilic drugs are readily dissolved although problems with absorption occur due to the hydrophobic conditions of cell membranes. This means that the cell permeation rate of hydrophilic drugs limits the overall bioavailability, due to the decrease in ability to diffuse across the cell membrane.1
Hydrophobic drugs are generally the opposite, where the dissolution rate and poor solubility therefore decreases the overall bioavailability. For oral formulation, the addition of a polar functional group onto the structure of a potential API is a proven way to enhance the aqueous solubility of hydrophobic drugs.4
Another common barrier for drug delivery is the immune system. Without specific targeting methods, the amount of drug that reaches its specified site is reduced, and can lead to concentrationg in the liver or spleen, potentially accumulating to toxic levels.5 In terms of biotherapeutics, siRNA molecules are degraded by 3’ mononuclease in blood plasma or by triggering an immune response. The largest challenge between siRNA’s and their potential application is an inherent negative charge, large molecular weight and hydrophilicity, in turn, reducing their ability to diffuse across cell membranes (∼13kDa).6
1.2. Nanoparticles to enhance drug delivery
The field of nanoparticle delivery systems has recently exploded in research popularity. The benefits of nanoparticle drug delivery systems is overall versatility, the ability to control release and specific targeting. Nanoparticles such as liposomes, niosomes, polymeric micelles and polymersomes have all been studied extensively, and have shown to have their own individual benefits whether its, cell permeation, loading capacity, stability, membrane versatility and also economic implications of each.5 Some liposomal delivery systems are currently FDA approved and are known to reduce toxicity to healthy cells with enhanced therapeutic effect compared to the administration of free drug.7 Liposomes however have a restricted loading capacity, and high concentrations can lead to a rupture in the membrane bilayer.5 Polymersomes however have higher loading capacity and stability, making them an exciting candidate in nano-drug delivery systems.
1.2.1. Polymersomes
Polymersomes have acquired substantial interest in recent years, as an answer to the issues with specificity, efficacy and controlling the kinetics of drug release.8 They are amphiphilic in nature and are formulated from hydrophilic-hydrophobic block copolymers, forming a lipophilic bilayered system with a hydrophilic core and hydrophilic outer membrane.9
Figure 1.2. Schematic Representation of Polymersome formation10
Polymersomes can be used as singular or multimodal targeted delivery systems and have huge versatility in their loading capacity. 11 The benefit of polymersomes is the ability to load hydrophobic and hydrophilic drugs, as well as the ability to load imaging agents and the encapsulation of other nanoparticles.8 The most distinctive feature of polymersomes however is the aqueous hydrophilic core.9 The ability to deliver therapeutics and diagnostics, compartmentalised into one “package” is particularly desirable in the field of theranostics. However, like any new drug or formulation, polymersomes and other biomimetic nanoparticles could encounter limitations in either early clinical trials or phase IV trials.
Research in nanoparticle stability has highlighted that polymersomes formulated from amphiphilic block co-polymers containing poly (ethylene glycol)(PEG) are more stable than other nanoparticles in systemic circulation and therefore sustain a longer time of residency.11, 12 This property has been attributed to the hydrophilic block copolymer PEG12 and also the thickness of the hydrophobic bilayer.8 The long PEG polymer chains on the outer hydrophilic membrane, forms a PEG brush which lowers immuno-sensitivity by resisting protein adsorption, allowing them to remain undetected in circulation.13 The thickness of the bilayer is determined by the length of the hydrophobic chains and therefore longer chains form thicker bi-layered membranes, leading to better protection of the encapsulated contents.
The administration of foreign biomolecules or free drugs can cause an immune response and are therefore engulfed by the mononuclear phagocytic system or broken down by enzyme degradation.11.13 However, the aqueous core of polymersomes, provides a bio-friendly environment for peptides, proteins, DNA and siRNA’s, which seems to be the focus for the future of personalised medicine. DNA and siRNA’s can be intravenously administrated but for biotherapeutic strategies to be effective they require extensive cellular uptake. 14 Potentially, this could be overcome by loading biomolecules into polymersomes with the attachment of a membrane penetrating peptide or through the exploitation of receptor-mediated endocytosis.14
1.2.2 Polymersome loading and membrane functionality
The versatility of polymersome membranes, enhances targeting abilities, therefore increasing therapeutic effect and inversely lowers the toxic side effects to healthy cells.12 Responsive or “smart” polymersomes formulated with “triggers” can cause polymersome degradation or pore formation, therefore releasing the encapsulated contents. “Smart” polymersomes can use pH, temperature, light, magnetic field and also multifunctional triggers which allows drugs to be directed and released in extremely precise locations.11
Loading techniques of polymersomes can be either remote or passive. Compounds with low water solubility use active loading procedures to achieve a high loading capacity. Passive loading involves the dispersion of the polymer in an organic solvent and the hydrophilic compound to be encapsulated is added at a later preparation stage.15 Encapsulation efficiency refers to the drug loading capacity of nano-drug delivery systems. Although lower encapsulation efficiency, results in the administration of more polymersomes to achieve a sufficient therapeutic effect leading to a higher amount of polymer in the body.16
1.3 Nanoparticle and ophthalmic drug delivery
Drug absorption is particularly difficult in regards to topical ophthalmic preparations, where the bioavailability of the active drug is around 1-2%.17 The reason for this comes down to the physiological limitations of the eye. It’s suggested that in order to increase the bioavailability of ocular topical formulations, they must possess sustained drug release, extended corneal contact time and satisfactory pre-corneal penetration.17 18
A study by Kasdorf 19 showed, that the permeability of the mammalian vitreous humor of the eye is highly dependent on the electrostatic interactions between the biopolymer network and diffusing particles. The research also shows that the penetrability behaviour of charged and neutral compounds applies more so to larger compounds (150kDa) and was statistically insignificant for smaller compounds (4kDa).19
1.4 Rationale for Research
It’s apparent that there is room for improvement in ocular drug delivery methods. Intravitreal injections can overcome many of the physical barriers of the eye, but with limited patient incompliance (up to 40%), mostly due to the fear of intravitreal injection.20 The introduction of nanoparticles has thoroughly enhanced the world of drug delivery, with increased cell permeation, increased solubility and targeted delivery. Liposomes are a great nano-drug carrier but appear to have limitations compared to polymersomes such as reduced stability and limited loading capacity.
Polymersomal drug delivery systems can be tuned to overcome the barriers presented at complex sites of action. Permeability of the vitreous humor is particularly difficult but possibility lies in the exploitation of electrostatic interactions i.e charged polymersomes. siRNA’s are unable to permeate cell membranes due to their net negative charge but could potentially be masked by encapsulation into a polymersome.
This research investigates the effect of charge on polymersomal drug delivery systems, in terms of encapsulation efficiency and release of neutral and negatively charged compounds.
2.0. Hypothesis and Aims
2.1. Hypothesis
The effect of a positive charge on the polymersome will have a direct influence in the ability to encapsulate and release negatively charged compounds. The electrostatic interactions between the opposite charge on the encapsulated compound will decrease the rate of drug release from the polymersome and also improve upon the encapsulation efficiency.
2.2. Overall Aim
To investigate the effect of positive charge of a polymersome in terms of characterisation and release profile of compounds with both neutral and negative charge.
2.3. Specific Aims
2.3.1. Prepare blank polymersomes using a reverse phase evaporation method. Dynamic light scattering (DLS) will be used to characterise the polymersomes, investigating the polydispersity index (PDI), size (nm) and zeta potential (mV) of the polymersomes.
2.3.2. Encapsulate FITC-Dextran or FITC-CM Dextran into the polymersomes. Analysis of, PDI, size (nm) and zeta potential (mV) of the now “full” polymersomes.
2.3.3. Analyse the encapsulation efficiency and release profile of fluorophore from positively charged and neutral polymersomes over a 24 hour period.
2.3.4. Compare release profile data, PDI, size and zeta potential, of neutral polymersomes and positively charged polymersomes.
3.0 Methods and materials
FITC-CM dextran powder, FITC- dextran powder, Sodium chloride, Sodium Bicarbonate, Calcium chloride di-hydrate, Phosphate buffer saline tablets, Dialysis tubing (Molecular Weight Cut Off(MWCO) 4000kDa), 5ml round bottom flasks (RBF), 50ml centrifuge tubes and 1ml Eppendorf tubes all sourced from Sigma Aldrich.
- Malvern Zetasizer Nano ZS
- Stuart™ hotplate stirrer SB162-3
- Branson 3510-DTH Ultrasonic Cleaner
- Büchi® Rotavapor® R-210 evaporator
- Büchi® vacuum pump V-700
- Buchi B-491 Heating Bath
- 19776 Angle Rotor
- ZEN1002 Universal Dip Cell
- DTS 1060 Capillary Cell
The polymers used in this project are referred to as P13 and P33+. P13 is a neutral polymer with an overall neutral charge. P33+ is a positively charged polymer with an overall net charge of 20+ mV. Both polymers were made in house and gifted by Ulster University.
Figure 3.1 Chemical Structures of P13 and P+33
3.1. Preparation of Stock solutions for Polymersomes
Stock solutions were prepared in 10ml vials of P13 in Chloroform (5mg/ml) and P13 in PBS (5mg/ml). Stock solutions of FITC Dextran and FITC-CM dextran were prepared with distilled water in 10ml vials at a final concentration of 2mg/ml and protected from light with foil. This process was repeated for the positively charged polymer, were P13 was replaced with P+33. A stock solution of P+33 in H2O with a concentration of (5mg/ml) was also prepared.
3.2. Formulation of Polymersomes
A reverse evaporation method was combined with further sonication to formulate the polymersomes. 0.5ml of P13 in Chloroform (5mg/ml) was added to a 5ml round bottom flask (RBF) and evaporated to dryness using a Büchi® Rotavapor® R-210 evaporator. 100l of FITC dextran (2mg/ml) was added to the RBF and evaporated to dryness. 1ml of chloroform was then added to the RBF and sonicated in a Branson 3510-DTH ultrasonic cleaner for 15 minutes. After sonication, 0.5ml of P13 in PBS (2mg/ml) and 0.5ml of PBS was added to the RBF and further sonicated for 30 minutes. The RBF was then placed back on to the rotavapor to remove any remaining solvents from the RBF.
To formulate the polymersomes, the same method was repeated replacing the according solutions where applicable, see Table 1.
Table 1. Components used in the creation of different polymersomes
- Polymersome *in chloroform (5mg/ml) Fluorophore (2mg/ml) *in PBS (5mg/ml) & PBS * in H2O (5mg/ml) & H2O
- Neutral Polymersome (Blank) P13 N/A P13 N/A
- Positive Polymersome (Blank) P+33 N/A P+33 N/A
- Neutral Polymersomes w/ FITC-Dextran P13 FITC Dextran P13 N/A
- Neutral Polymersomes w/ FITC-CM Dextran P13 FITC-CM Dextran P13 N/A
- Positive Polymersomes w/ FITC Dextran (H2O) P+33 FITC Dextran N/A P+33
- Positive Polymersomes w/ FITC-CM Dextran (H2O) P+33 FITC-CM Dextran N/A P+33
- Positive Polymersomes w/ FITC-CM Dextran (PBS) P+33 FITC-CM Dextran P+33 N/A
3.3 Characterisation of Polymersomes, Polydispersity Index and Size
Dynamic light scattering (DLS) was used to characterise the polydispersity index and size of the polymersomes, this was performed in a Malvern Zetasizer Nano ZS. Using a pipette, 100l of polymersomes was transferred from the RBF into a 1cm glass cuvette and diluted with 900l of H2O to make 1ml of a diluted polymersome solution. The solution was mixed using the pipette tip and the glass cuvette placed into the zetasizer. Analysis parameters were set at a temperature of 25C and results were recorded on the software.
3.3.1. Zeta potential of Polymersomes
1ml of diluted polymersomes was transferred from the glass cuvette to a folded capillary cell (DTS1060). The DTS1060 was placed into the zetasizer and the zeta potential analysis was run at a temperature of 25C.
The DTS1060 was replaced with a ZEN1002 dip cell, for the positively charged polymersomes, as the zeta potential fluctuated excessively between runs of the same solution. After PDI and Size analysis, positive polymersomes were then transferred from the glass cuvette to a plastic cuvette. The ZEN 1002 universal dip cell was inserted into the plastic cuvette and zeta potential analysis was run at 25C.
3.4. Formulation of Simulated Tear Fluid solution
500 ml of simulated tear fluid solution was formulated with the composition.18
Sodium Bicarbonate – 0.2% w/w
Calcium Chloride di-hydrate – 0.008% w/w
Sodium Chloride – 0.67% w/w
The components were weighed on a microbalance in separate weigh boats, transferred to a 1 litre measuring cylinder and then made up to 500ml with deionised water. The contents of the measuring cylinder were then poured into a 1 litre glass jar.
3.5. Release study
Dialysis tubing was cut into three pieces of around 5cm and placed into a beaker of water overnight. The molecular weight cut off (MWCO) of the dialysis tubing was 4000kda, which allows the FITC-dextran (4000 Kda) to outflow from the dialysis bags. The bags were then triple tied at one end. 500l of polymersomes were pipetted into each dialysis bag and triple tied at the other end. Each bag was then checked for leakage. The tied bags were then locked into the top of individual 50 ml centrifuge tubes by hanging the loose thread over the lip of the lid. The 50ml centrifuge tubes were then inserted into a 19776 angle rotor and centrifuged in a Sigma 3-30 K centrifuge for 2 hours at an RCF of 3000 and a temperature of 4C.
Three 30ml vials were filled with 10ml of simulated tear fluid and a magnetic stirrer put in each vial. The vials were then wrapped with tinfoil and placed on a Stuart™ hotplate stirrer SB162-3. The temperature of the simulated tear fluid was adjusted to 37 2C. A beaker of around 40ml of simulated tear fluid was left beside the hotplate stirrer for ease of access. The centrifuge tubes were removed from the centrifuge and wrapped in foil to avoid contact with light. The dialysis bags were then removed from the centrifuge tubes and each bag was placed into a separate vial, which was also wrapped in foil. A stop-clock was started after placing the bags in each of the vials and the lids were placed onto the vials.
1ml samples from each jar were removed at time intervals of 0.25, 0.5, 1, 2, 4 , 8 and 24 hours and stored in 1ml Eppendorf tubes and again protected from light. After 1ml of sample was taken from the vials, the volume was returned to 10ml by pipetting 1ml of simulated tear fluid into each vial.
The samples from the release study were analysed using a Varian Cary Eclipse Fluorimeter. The 1ml samples were removed from the Eppendorf tubes and pipetted into a 1cm glass cuvette. The glass cuvette was then placed into the fluorimeter and the excitation wavelength was set to 490 nm with an emission detection range between 500 – 700 nm. This process was repeated for each sample, washing between each reading. The peak values were then recorded from the graphs.
3.6. Encapsulation efficiency
After centrifugation, any un-encapsulated fluorophore fell to the bottom of the centrifuge tubes. The remaining volume left in tube was then measured using a micropipette. This was then pipetted into a 1cm glass cuvette and made up to 1 ml with simulated tear fluid. The glass cuvette was then placed into a Varian Cary Eclipse Fluorimeter. Excitation wavelength was set at 490 nm, slit width of 5nm and emission range was set from 500-700nm. From the resulting graph the peak value was recorded.
3.7 Standard curves of Fluorophores
0.25ml of FITC-Dextran (2mg/ml) was pipetted into a 25ml volumetric flask and made up to the mark with simulated tear fluid to make a final concentration of 20g/ml solution (Stock 2). Serial dilutions of, 2, 4, 8, 12, 16g/ml were prepared from stock 2 and made up with simulated tear fluid in 10ml volumetric flasks. 1ml from each volumetric flask was then pipetted into a 1cm glass cuvette and analysed on a Varian Cary Eclipse Fluorimeter. The excitation was set to 490nm with an emission detection range of 500-700nm with a slit width of 5nm. Results from the fluorimeter graphs, were then plotted on a graph of concentration (g/ml)/ fluorescence (a.u), were the equation of the line and R2 value were devised.
The method above was repeated for FITC-CM dextran.
4.0 Results & Discussion
4.1. Preparation and characterisation of blank polymersomes
Polymersomes were successfully prepared as described in section 3.2. They were confirmed using DLS. All polymersomes were found to be within the anticipated size range of less than 500 nm, which is essential for endocytosis to occur. The results of the neutral polymersome prepared from P13 and that of the positively charged polymersome, prepared form P+33 are shown in Figure 4.1.
Figure 4.1 Size of empty polymersomes
The size of both polymersomes were found to be similar, and both below 200nm. The positively charged polymersome is slightly smaller, this may be indicative of a smaller hydrophobic bilayer, but further analysis would be required in order to confirm.
The PDI is used as a measure to determine how monodisperse the colloidal suspension is, the results of the two polymersomes prepared are shown in Figure 4.2.
Figure 4.2 Polydispersity index of empty polymersomes
The PDI of both polymersomes prepared are below 0.5, and therefore indicative of a monodisperse system. The neutral Ps had a slightly better PDI of 0.244, however it did not appear to be a significant difference between the two polymersomes prepared.
Finally, characterisation of the two polymersomes was confirmed using the zeta potential. As can be seen from Figure 4.3, the surface charge of the polymersome prepared from the neutral polymer was negligible, however that of the positively charged polymersomes was +22.7 mV. Although these polymersomes were loaded, FITC dextran is a neutral compound and therefore did not affect the zeta potential of the polymersomes.
Figure 4.3 Zeta potential of polymersomes
The above results confirms that both polymer 13 and polymer +33 can be formulated into polymersomal structures suitable for drug delivery.
4.2. Characterisation of Polymersomes loaded with fluorophores
FITC-dextran was chosen as a biological mimic, with FITC-CM dextran as a negatively charged analogue, the structure of FITC-CM dextran is shown below in Figure 4.4.
Figure 4.4 Structure of FITC-CM dextran
The carboxyl groups highlighted in Figure 4.4 gives the FITC-CM dextran its negative charge. These groups are therefore absent in FITC-Dextran.
4.2.1. Characterisation of loaded Polymersomes size
FITC-CM dextran and FITC dextran were both successfully encapsulated into neutral and positively charged polymersomes as described in Section 3.2 and again confirmed using DLS.
Figure 4.5 Size of loaded neutral polymersomes
Figure 4.5 confirms that the neutral polymersomes prepared from P13 remained smaller than 500nm. There was a small but insignificant increase in loaded polymersome size compared to the unloaded polymersomes, and assumed this is just an effect of loading. Polymersomes loaded with FITC-CM dextran had a slightly larger size than FITC-Dextran, this could be due to electrostatic repulsion between FITC-CM dextran molecules (see Fig 4.4), therefore increasing the size of the polymersomes.
Figure 4.6 Size of loaded positive polymersomes
Results of the loaded positive polymersomes in Figure 4.6 showed again that polymersome size had increased after loading of the fluorophores. Positive polymersomes loaded with FITC-CM dextran were almost double in size of those with FITC Dextran and exceeded the ideal 200nm size. This would suggest that the electrostatic attraction between polymersome and fluorophore increased the amount encapsulated in the aqueous core. It could also be due to excess binding of FITC-CM dextran to the outside of the polymersome membrane due to the same electrostatic attraction. Further research to confirm the location would be ideal, possibly using bioimaging.
4.2.2. Characterisation of Polymersomes using PDI
After encapsulation of the fluorophore, the PDI of loaded neutral polymersomes increased and the systems became more poly-dispersed as seen in Figure 4.7. The PDI of FITC-dextran polymersomes just exceeded over 0.5 which is on the boundary of acceptability for nanoparticle dispersion system. A PDI of over 0.5, means that the system is more poly-dispersed. It may be suggested that a longer sonication time for loaded polymersomes may be ideal to achieve a more monodispersed system, compared to the sonication times outlined in 3.2.
Figure 4.7 Polydispersity Index of Neutral Polymersomes
Figure 4.8 Polydispersity index of Positive Polymersomes
Figure 4.8, shows that there was no significant difference in PDI between loaded and unloaded polymersomes. In contrast to the results of Figure 4.7, the positive charge of the polymersome may be a contributing factor to uniformity, therefore resulting in a more monodispersed system. The PDI of dispersion systems is lowered by charged particles but to confirm this was in true effect, polymersomes with differing charges should be tested.
4.2.3. Characterisation of Polymersomes using zeta potential
Figure 4.9 Zeta Potential of loaded Neutral Polymersomes
There was no significant difference in zeta potential between both loaded neutral polymersomes. The results from Figure 4.9, does suggest that the charge of FITC-CM dextran becomes undetectable at the surface membrane of neutral polymersomes. After release studies it would be ideal to test the zeta potential of the released FITC-CM Dextran to confirm it still possessed a negative charge.
Unlike Figure 4.9, there is a clear significant difference between the zeta potential of the positive polymersomes in Figure 4.10. It appears that the negative charge of FITC-CM reduces the zeta potential of the positive polymersome. It could be suggested that in the formulation (section 3.2), addition of negatively charged species somewhat neutralizes the positive charge on the polymer. It could also be to accumulation of FITC-CM dextran molecules on the surface of the polymersome membrane as mentioned in Section 4.2.1.
The results in Table 2. confirm the charge on P5 and P3. P4 however shows a negligible zeta potential. Comparison of P2 and P3 shows that FITC-CM dextran lowers the zeta potential in positive polymersomes (H2O). Although comparison of P1 and P4 shows that if PBS is used in the formulation of positive polymersomes zeta potential is reduced, not because of FITC-CM dextran but due to formulation with PBS.
Unfortunately this was only discovered part way through the research project
therefore future work should include formulating the neutral polymersomes with H2O, to give a fair comparative result.
Table 2. Zeta potential of Polymersomes formulated with PBS and H2O
Polymersome Polymersome/Polymer Zeta potential (mV)
- P1 Positive Polymersome (PBS) with FITC-CM dextran -0.04 0.2
- P2 Positive Polymersome (H2O) with FITC-CM dextran 3.5 1.3
- P3 Positive Polymersome (H2O) with FITC dextran 22.7 2.9
- P4 Blank Positive Polymersome (PBS) 0.22 0.3
- P5 P33+ in water 20.6 1.7
4.3. Release and Encapsulation studies
The release and encapsulation studies were carried out as per section 3.5, 3.6 and 3.7. The equations derived from Figure 4.11 and Figure 4.12 were used to calculate the concentration of fluorophore in the samples.
4.3.1. Standard Calibration of encapsulated compounds
Figure 4.11 Calibration curve of FITC dextran (excitation wavelength 490nm)
Figure 4.11 shows the results from section 3.5, the method was performed in triplicate and an average taken. Error bars displayed are the standard deviation for each point. The equation of the line for FITC dextran(R2 = 0.9981):
Figure 4.12 Calibration curve of FITC-CM dextran (excitation wavelength 490nm)
Figure 4.12 shows the results from section 3.5. The equation of the line for FITC-CM dextran was (R2 = 0.9978):
R2 values were 0.9978 and 0.9981 respectively, indicating strong linearity between the concentration of fluorophore and fluorescence (au). The equations derived from Figure 4.11 and Figure 4.12 were also used in sections 4.3.2 and 4.3.3.
4.3.2. Encapsulation efficiency
Encapsulation efficiency determines how much of the fluorophore was encapsulated in the polymersomes. In reference to section 3.6, 100l of fluorophore was loaded into each polymersome. After centrifugation any un-encapsulated fluorophore in the bottom of the centrifuge tubes was made up to 1ml and analysed in the fluorimeter with a 1cm glass cuvette. Using the equation of the line the concentration was calculated and then subtracted from the original loaded 100l of fluorophore. The difference is then worked out as a percentage and results displayed in Table 3.
Table 3. Encapsulation efficiency of Polymersomes
Polymersome Encapsulation Efficiency %
- Neutral Polymersome (PBS) (FITC) 64.55 5.75
- Neutral Polymersome (PBS) FCM) 71.75 1.6
- Positive Polymersome (H2O) (FITC) 84.83 9.72
- Positive Polymersome (H2O) (FCM) 93.12 0.14
Table 3. shows the encapsulation efficiency of the neutral polymersomes formed from P13, and the results were fairly close to previous research using the same polymer.10 Neutral polymersomes encapsulating FITC-CM dextran had slightly higher encapsulation efficiency (71.75 1.6), but the standard deviation from neutral polymersome with FITC shows it was not of any significance.
The greatest significance however was the encapsulation efficiency of the positive polymersomes overall. However both positive polymersomes had higher encapsulation efficiency, which indicates that the P+33 polymer structure is a contributing factor, rather than the attraction between positive polymersomes and negative compounds.
4.3.3. Release of FITC-CM and FITC dextran from Polymersomes
Figure 4.15 Release of FITC dextran & FITC-CM dextran from Neutral Polymersomes
Release profile of neutral polymersomes is shown in Figure 4.15. There is a clear significant difference between the amount of FITC-CM dextran and FITC dextran released from the neutral polymersomes. This could be attributed to the ionic strength of the simulated tear fluid, although to confirm whether or not this is a true effect of ionized solutions, further release studies should be also carried out in deionised water.
Figure 4.16 Release of FITC dextran & FITC-CM dextran from Positive Polymersomes
Positive polymersomes release studies (Figure 4.16) showed that the amount of fluorophore released over time was generally lower than the results of Figure 4.15. There wasn’t any significant difference between the release profile of positive polymersomes loaded with FITC-CM dextran or FITC dextran. This suggests that the difference of charge between loading compounds is negligible in relation to the amount and rate of release from positive polymersomes.
Figure 4.17 Release of FITC CM dextran from Neutral and Positive Polymersome
Figure 4.17 compares release profiles of FITC-CM dextran of neutral and “positive” polymersomes, even though the positive charge was lost as mentioned in 4.2.3. It appears that the polymersomes formulated from P+33 released more FITC-CM dextran than the P13 polymersomes, over a 24 hour period. The results in Figure 4.17. were highly unexpected, however, it could be due to steric or electrostatic repulsion between corona blocks of P+33 polymer.21 Its also quite possible that not all of the free FITC-CM dextran was removed after centrifugation and remained attached to outer membrane of the positive polymersomes. This would suggest that length of centrifugation time could be extended.
Figure 4.18 Release of FITC-CM dextran from positive polymersomes, PBS & H2O.
The effect of using PBS compared to H2O in section 3.2 is shown in Figure 4.18 above. The release of FITC-CM from Poly+(H2O) is significantly lower than Poly+(PBS). This confirms that the charge on Poly+(H2O) reduces the amount of negatively charged species release, as Poly+(PBS) was confirmed to have lost charge in Table 2.
4.4. Comparison of data
Comparison of positive and negative polymersome size showed that encapsulating FITC-CM dextran increased the size of the polymersomes. Positive polymersomes however drastically increased in size and was decided this was a direct effect of the charge attraction between positive polymersomes and the negative species. The PDI of neutral polymersomes increased after encapsulation of both fluorophores, whereas, the positive polymersomes remained fairly monodispersed, before and after encapsulation. This is due to the positive charge of the polymersome increasing the stability of the dispersion system.
There was no difference between the zeta potential of loaded neutral polymersomes, suggesting that the negative charge of FITC-CM was masked, although this was not fully confirmed. In this case release studies should be performed in water and the zeta potential of FITC-CM should be analysed from the 1ml samples taken after release. By using water in the release instead, will also protect the integrity of the method as any ionic interactions form the simulated tear fluid caused false readings with the dip cell.
There was a direct effect of FITC-CM dextran on the zeta potential of positive polymersomes. The zeta potential was reduced and it was confirmed that this was due to FITC-CM dextran.
It appeared that the zeta potential was reduced of polymersomes formulated with P+33 and PBS, although after replacing PBS with H2O, the zeta potential was still reduced after encapsulation of FITC-CM dextran. Encapsulation of fluorophores in positive polymersomes was higher than neutral polymersomes. Although encapsulation efficiency was higher it cannot be confirmed that this was a direct effect of charge attraction, and more so an effect of using H2O in formulation of the polymersomes.
The release studies showed that positive polymersomes formulated with H2O released less fluorophore over a 24 hr period. However the charge attraction between positive polymersome and FITC-CM dextran was not the reason for this as, positive polymersomes with FITC- dextran showed a very similar profile over 24 hours. PBS is used as a buffer to regulated pH, which may be the contributing factor to the release profiles of positive polymersomes, future work should include regulation of pH in release studies as this was not tested in the method. Release profiles of neutral polymersomes showed that the rate of release was determined by interactions with ionized solutions and therefore should be repeated in H2O to determine if this was truly the contributing factor.
5.0 Conclusion
Biotherapeutic strategies are growing and the ideology of personalised medicine is slowly building momentum.The delivery of biomolecules such as siRNA come with various issues, one of which, is an inherent negative charge. This property then limits siRNA’s ability to permeate cell membranes, which also have a net negative charge.
The versatility of polymersomes makes them an exciting candidate for a multitude of application and also provides a bio-friendly aqueous core suitable for the likes of siRNA’s. By investigating the effects of charge on polymersomal delivery systems, FITC-CM dextran was used as a biological mimic of siRNA and this research provides preparative data for future work applicable to ocular drug delivery and siRNA delivery.
Polymersomes using P13 and P+33 were successfully formulated using a reverse phase evaporation technique and characterised using DLS ((P13 polymersomes, PDI: 0.244 0.08, HDR: 169.4 21.2 nm)(P+33 Polymersomes, PDI: 0.386 0.084, HDR: 126.6 38.4)). No data was recorded for zeta potential of unloaded polymersomes.
FITC-dextran and FITC-CM dextran was loaded successfully into both neutral and positive polymersomes and characterised using DLS. Characterisation of PDI, Size and ZP (see. Appendix).
From the data the hypothesis can only be partly accepted. Positively charged polymersomes had a direct influence on encapsulation efficiency of negatively charged compounds (Table 3. P+(FITC-CM), 93.12 0.14%). Although the rate of release was lower for positive polymersomes, similar release profiles were seen for negative and neutral fluorophores, therefore it was not a direct effect of charge attraction (Figure 4.16).
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