Research article Received: 5 June 2014,
Revised: 13 August 2014,
Accepted: 4 September 2014
Published online in Wiley Online Library
(wileyonlinelibrary.com) DOI 10.1002/bio.2789
Transfecting pDNA to E. coli DH5α using bovine serum albumin nanoparticles as a delivery vehicle Jitendra Wagh,b Kuldeep J. Patel,c Parth Soni,c Krutika Desai,c Pratik Upadhyayd and Hemant P. Sonia* ABSTRACT: We describe the formulation of bovine serum albumin nanoparticles (BSA-NPs) by the coacervation method using surfactants. Plasmids (pUC18, pUC18egfp and pBBR1MCS-2) isolated from E. coli were incorporated into the BSA matrix by incubating in albumin solution prior to formulation of NPs. Plasmid incorporation was calculated by % yield, entrapment efficiency, DNA loading capacity and release of entrapped DNA by comparing with blank NPs. BSA-DNA binding studies were carried out by using fluorescence spectroscopy and Fourier Transform Infra Red Spectroscopy (FT-IR). The surface charge distribution of the NPs loaded with plasmid was calculated using zeta potential. The photoluminescence of BSA-NPs was quenched when loaded with pDNA, confirming the interaction of DNA with BSA. Altogether, these results provide evidences for the excellent DNA carrying efficiency of BSA-NPs without loss of plasmid’s integrity. The NPs were used to transfect E. coli DH5α strain lacking ampicillin resistance. They, however, showed ampicillin resistance subsequent to transfection with plasmid encoding ampicillin resistance gene. Effect of transfection was confirmed by confocal microscopy and by the isolation of the plasmid by agarose gel electrophoresis from the transfected bacterial culture. This study clearly demonstrates the efficacy of BSA-NPs as delivery vehicle for pDNA transfection. Copyright © 2014 John Wiley & Sons, Ltd. Additional supporting information may be found in the online version of this article at the publisher’s web site. Keywords: BSA nanoparticles; Gene delivery; Enhanced green fluorescence protein
Introduction Transfection of foreign plasmid DNA or RNA into host cell nucleus to modify, change, or silence expression of gene is a challenging task. Availability of strategy in this context can ensure the production of specific protein that can help to mitigate conditions like Parkinson’s disease (1) and cystic fibrosis (2) or even provide relief to painful chemotherapy treatments in cancer (3). The cellular nuclease enzymes constitute the major obstacle in this task as they can degrade naked plasmids in the cytoplasm before entry into the nucleus leading to low transfection efficiency (4–6). Various strategies have been devised to overcome this problem like trapping such pDNA in a specialized carrier of both natural and synthetic origin (7). Such a carrier should not only bind with pDNA but also with receptors on the cell membrane for successful entry into cells. After entering cytoplasm, it should neither interact with other organelles nor interfere with any biological process (8). Ideally, it should be degraded once it has entered the cytoplasm and release the cargo without causing any harm to the cell. There are carriers that can directly traverse the cell membrane and enter the cytoplasm and viruses are the naturally evolved machinery ideal for such a process (9,10). However, there are serious drawbacks with such systems like with immunogenicity (11,12), oncogenicity (13) and recombination efficiency (14), which restrict the successful application of them. Many chemical agents such as cationic lipids like 2,3-di-oleoyloxytrimethyl ammonium propane (15), fullerenes and their derivatives (16,17), biodegradable polymers and dendrimer based materials e.g. PLL-b-PEG (18–20), PAGA (21), PAMAM type dendrimers etc.; carbohydrate based polymeric cages e.g. β-cyclodextrins (22,23), chitosan (24,25) etc. have been
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considered as nonviral gene-delivery vectors. But they also have their own drawbacks, e.g. high concentrations of fullerene-based materials can induce inflammation promoting the development of cancer (26). Recent reviews have focused on different types of gene-delivery systems, barriers encountered during nonviral gene delivery and the techniques to overcome them (27,28). Considering these, there is a need for development of efficient and specific delivery vehicles either natural or synthetic. Protein, being a macromolecule, can very well satisfy all such requirements to act as a gene-delivery vector (29). They are capable of selfassembly into various shapes and morphologies with well defined
* Correspondence to: Dr. Hemant P Soni. Department of Chemistry, Faculty of Science, The Maharaja Sayajirao University of Baroda, Vadodara-390 002, Gujarat, India. E-mail: [email protected] a
Department of Chemistry, Faculty of Science, The Maharaja Sayajirao University of Baroda, Vadodara-390 002, Gujarat, India
b
Department of Biochemistry, Faculty of Science, The Maharaja Sayajirao University of Baroda, Vadodara-390 002, Gujarat, India
c
Department of Biotechnology, Ashok & Rita Patel Institute of Integrated Study & Research in Biotechnology and Allied Sciences (ARIBAS), Sardar Patel University, V V Nagar, Gujarat, India
d
LJ Institute of Pharmacy, Sarkhej, Ahmedabad, Gujarat, India Abbreviations: AAV, adeno-associated virus; BSA, bovine serum albumin; BSA-NPs, bovine serum albumin nanoparticles; DLS, Dynamic Light Scattering; ESGT, European Society of Gene Therapy; FTIR, Fourier transform infra red; LAFU, laminar air flow unit; LB, Luria Broth.
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J. Wagh et al. pockets. For example, ferritin, a natural protein, has a cavity of 12 nm outer diameter and 8 nm inner diameter (30), pDNA can easily be accommodated intact in such a cavity. They can also be degraded naturally once their function is finished without causing any side effects, even more, their disintegration can be controlled and made more site specific. Hence, proteins have the potential to function as safer, non-toxic, stable and biodegradable genedelivery vectors (31). The purpose of this study was to develop a novel transfection technique by using BSA-NPs. At the initial stage, to demonstrate the efficiency of the developed protein nanoparticles as vehicle, we selected bacterial cells as host to deliver pDNA. However, this strategy may be further extended to eukaryotic cells as well. It has been observed that bacteria can take up protein nanoparticles as a part of their nutrient requirements (32,33). This can therefore, be exploited to transfer biomolecules like pDNA into a bacterium. The entrapment of pDNA in such protein nanoparticles may be an effective means to transfect the gene of interest into a host bacterium. Three different plasmids pUC18 (34), pUC18egfp (35) and pBBR1MCS2 (36) encoding kanamycin and ampicillin resistance genes, were isolated from E. coli culture. These plasmids were eluted, purified and utilized for loading on BSA-NPs. Such gene loaded BSA-NPs were then transfected into E. coli DH5α. As an evidence of successful transfection, bacteria under study were tested for the expected resistance against the specific antibiotics (37). Enhanced green fluorescence protein tag was attached with pUC18 as marker and confocal microscopy of the pUC18egfp transfected E. coli DH5α strain was carried out to confirm successful transfection. In this work, we have presented direct evidence of cell transfection by plasmid-loaded BSA nanoparticles.
Formulation of nanoparticles by coacervation method BSA (2% w/v) solution was prepared in water at 5.5 pH and added to pDNA (Table S2) and incubated for 1 h at room temperature. Then, the aqueous phase was dissolved in chilled ethanol drop wise (2:1, ethanol: water) at a rate of 1 ml/min. The coaservates, so formed were hardened with 25% glutaraldehyde solution (1.56 μg/mg) and further incubated for 2 h. The resulting nanoparticles were purified by centrifugation at 15,000 rpm for 10 min at 4 °C. The supernatant obtained was transferred in another microcentrifuge tube to estimate the amount of protein and unloaded DNA. The resultant pellet of purified nanoparticles was dried and stored at –20 °C. Transfection of E. coli DH5α with plasmid-loaded BSA nanoparticles For the uptake of nanoparticles by E. coli DH5α, pure culture (50 μL) was inoculated in LB (1 g/100 ml instead of 2 g/100 ml) containing filtered NPs solution (200 μL), under LAFU. The culture was allowed to grow over night at 37 °C with constant shaking (Table S3 and S4).
Experimental Materials Potassium acetate, glacial acetic acid, phenol, chloroform, sodium dodecyl sulphate (SDS), 25% glutaraldehyde were purchased from Merck. The restriction enzyme EcoRI, ampicillin, kanamycin, 10× assay buffers and other required chemicals were purchased from the Genei Chemicals, India. Plasmids, pBBR1MCS-2 (a plasmid originally derived from the pBBR1 plasmid of Bordetella bronchiseptica) express kanamycin resistance while, pUC18 and pUC18egfp show ampicillin resistance, which are high copy number plasmids (Table S1). Characterization of pDNA The bacterial cells were harvested to obtain the culture for isolation of plasmid DNA. Single colonies from pure culture were collected in a wire loop and inoculated in 100 ml autoclaved LB broth containing required antibiotic (5 μg/ml). The sample was incubated at 37 °C with 200 rpm overnight for uniform aeration and growth. Plasmid was isolated from bacteria by miniprep method (38). The isolated plasmid was detected by agarose gel electrophoresis (39). The observed bands of plasmid DNA were separated using a sharp sterilized blade, and added to a clean and dry microcentrifuge tube containing fixed weight of gel. In accordance with the weight of gel, three volumes of QG buffer was added. The tube was incubated at 50 °C for 10 min to dissolve the gel. To this, 1 volume of isopropanol was added and mixed thoroughly. The sample was now applied on to the column and centrifuged for 1 min, at 4 °C at 13,000 rpm. To this,
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750 μL of PE buffer was added and the column was allowed to stand for 3 min, followed by centrifugation for 1 min. The flow through was discarded and the column was placed in a clean microcentrifuge tube and allowed to air dry for 5 min. The eluted and purified pDNA was further used for restriction digestion using EcoRI, whose specific restriction sites are present in the plasmids. This helped in size determination of the isolated plasmids. For restriction digestion, the following cocktail was prepared for each sample of DNA consisting of template 50 μg, assay buffer (10×) 2 μL, BSA (10×) 2 μL and EcoRI (10 U/μL). In order to make up the total volume to 20 μL, autoclaved distilled water was added. These mixtures were incubated for 4 h at 37 °C and there after samples were run on 1% agarose gel.
Screening of transfectants To screen successful transfectants, the culture was aspirated (50 μL) and added to LB containing antibiotics for the plasmids used in study. This was allowed to grow over night at 37 °C. Successful growth of bacteria confirmed the uptake of plasmid from NPs with ampicillin resistance. This was further established by performing plasmid isolation using the culture which initially lacked any plasmid. For additional confirmation of successful transfection, bacterial cells were washed with normal saline for egfp expression present in pUC18egfp and then confocal microscopy (Carl Zeiss Microscopy) was carried out at excitation wavelength 485 and emission at 510 nm.
Characterization and evaluation Evaluation of pDNA The concentration and purity of plasmid DNA was measured in a UV spectrophotometer. For the purpose, pDNA solution (50 μL) was mixed in distilled water (2950 μL) and the absorption spectra were measured at 260 and 280 nm. The concentration of pDNA was expressed as ng/μL. The purity of DNA was assessed by 260/280 nm absorption ratio (Table S5). The pDNA was subjected to restriction digestion using EcoRI enzyme and agarose electrophoresis was carried out to determine the size
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Transfecting pDNA to E. coli of the eluted pDNA. The linear DNA fragments detected after digestion were separated on a 1% agarose gel along with the reference ladder.
pDNA added and the amount present in the supernatants obtained after the purification by spectrophotometer at 260 nm. DNA loading capacity was calculated as,
Evaluation of nanoparticles
% DNA loading capacity ¼
The mean size of formulated NPs was determined by Dynamic Light Scattering (DLS) technique. A BIC 90 plus (Brookhaven) equipped with 35.0 mW solid-state laser operating at 660 nm and an avalanche photodiode detector was used for the measurement of particle size distribution by DLS and surface charges were measured in term of zeta potential by a Zeta analyzer system, Nano Series (Malvern Instruments). The surface morphology and size of NPs formulations were also studied by Transmission Electron Microscopy operating at 200 kV (Philips CM200). The amount of protein transformed in NPs was determined by standard BCA protein assay. NPs obtained after centrifugation were digested in basic medium for 2 h at RT. The absorbance of the resulting solutions was measured at 562 nm. The% yield of NPs was calculated as, % yield ¼
Total weight of NPs 100 total weight of pDNA and BSA
(1)
%Entrapment efficiency ¼
Amount of DNA in NPsðμgÞ100 Yield of NPsðmgÞ (2)
Amount of pDNA present in NPs Amount of pDNA used in formulation
(3) Plasmid-loaded BSA-NPs were suspended in phosphate buffer saline (PBS, 300 μL) at pH 7.4 as release media in microcentrifuge tubes placed in shaking water bath at 37 °C at 60 rpm. At predetermined intervals, 50 μL samples were harvested and replaced with 50 μL fresh media. For the determination of amount of DNA released, the tubes were centrifuged and supernatant obtained was analyzed by using spectrophotometer at 260 nm. % Release of pDNA ¼
Amount of pDNA released Amount of pDNA present in NPs (4)
BSA-DNA binding study
Results and discussion
The interaction between BSA and pDNA in aqueous solution was investigated using fluorescence spectroscopy by means of Jasco FP-6300 spectrofluorometer using Xenon lamp as the excitation source. UV-Vis absorption spectra were recorded on a Perkin Elmer Lamda 35 uv/vis spectrometer. The binding constant Ksv was determined using Stern-Volmer equation (40). The required, λmax of BSA was measured by UV spectroscopy. This λmax value, in turn, was used to excite loaded and unloaded BSA nanoparticles for emission spectrum in fluorescence spectroscopy, by keeping the concentration of BSA constant, and simultaneously increasing the amount of pDNA. Loading of pDNA on BSA nanoparticles and interaction of pDNA with BSA nanoparticles were also studied. For the purpose, the absorption spectra of pure BSA, pure pDNA, BSADNA solution, blank nanoparticles (only BSA) and pDNA loaded nanoparticles, were obtained by using a UV spectrophotometer. Samples with appropriate concentration were scanned at 200–400 nm keeping water as blank. Fourier transform infrared (FTIR) spectra of the samples were recorded in the region of 500–4000 cm–1 using Perkin Elmer FTIR Spectrometer (Spectrum RX1).
The plasmids (pUC18, pBBR1MCS2 and pUC18egfp) were isolated from E. coli DH5α and loaded on BSA-NPs for cell transfection. An absorption ratio (260:280 nm) around 1.8 indicates that the isolated plasmids are pure and not contaminated by any protein impurity. Per cent yield, pDNA loading capacity and pDNA loaded on the NPs data suggest that BSA can be an ideal protein to serve as vehicle for pDNA transfection (Table S2,S3 and S4). TEM images show BSA-NPs to be spherical and monodispersed in the size range of 150–170 nm (Fig. 1). It can be observed that on binding pUC18 on BSA-NPs, the size increases in the range of 180–200 nm with almost no change in shape or dispersity. This is also true for other two plasmids as well. The DLS plots for free and bound BSA-NPs (100 vs. 200–300 nm) clearly support the TEM observation (Fig. S2). Zeta potential measurement for free NPs (–53 meV) indicates that the aqueous dispersion of NPs is quite stable. On pDNA binding, the value decreases to lower negative (Table 1) indicating depletion of charge and agglomeration (140–200 nm) of NPs (37). From the zeta potential values, the mechanism of interaction of BSA-NPs with pDNA may be predicted. While in the aqueous medium at acidic pH (5.5), the N and C terminals of the protein are positively and negatively charged respectively, the negative charge gets partially neutralized on pDNA binding due to which a marked depletion of charge occurs in the bound NPs. On comparing Tables 1 and S2 it can be seen that the size of the plasmid also matters for the depletion of charge. As the size increases, more charge on the carrier neutralizes resulting in depletion of charge of the overall protein-plasmid assembly. Fluorescence spectroscopy is an important tool to study the interaction among biomolecules and NPs. It can not only suggest the mechanism of interaction but also provide clues about the micro-environment of fluorophore (BSA in this case) from which fluorescence originates. It can also provide information
Evaluation of plasmid DNA loaded nanoparticles The absorbance of BSA solution with different concentrations in basic medium was measured by UV spectrophotometer at 562 nm using 0.1 M NaOH solution as blank. From the data, a calibration curve of concentration versus absorbance was plotted (Fig.S1). Further, standard calibration curve was obtained for eluted pDNA solution in water at 260 nm. Amount of pDNA released from nanoparticles can be estimated on comparing the curves obtained by UV spectrometry with the standard curve. The amount of pDNA loaded on BSA nanoparticles was calculated as the difference between the total amount of the initial
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Figure 1. TEM images of (a) unloaded (b) pUC18 (c) pBBR1MCS-2 and (d) pUC18egfp loaded BSA nanoparticles.
Table 1. Zeta potential of BSA nanoparticles loaded with different plasmids Nanoparticles Loaded with size of the Zeta potential plasmid plasmid in kb (mV) BSA BSA BSA BSA
– pUC18 pUC18egfp pBBR1MCS-2
– 2.9 3.5 5.1
–53.2 –41.6 –9.04. –10.4
about conformational changes taking place when protein interacts with other biomolecules (41). Tryptophan (Trp), tyrosine (Tyr) and phenylalanine (Phe) residues are mainly responsible for fluorescence in BSA. While one tryptophan residue (Trp134) is located in the first domain the other (Trp-212) is located in the second domain of BSA protein. Trp-212 is situated inside the hydrophobic binding pocket while TRP-134 is on the outer hydrophilic surface of the BSA (42). Fig. 2 shows the fluorescence emission spectra of BSA in presence of different concentrations of pUC18, pUC18egfp and pBBR1MCS-2 following an excitation at 280 nm. It can be seen from Fig. 2 that BSA fluoresces strongly at 675 nm when excited at 280 nm at pH 7.4 and its intensity gradually decreases with increasing concentration of pDNA. Further, the blue shifting in the λmax value occurs with increase in concentration. When pDNA comes in the vicinity of the protein, there are two possibilities: (i) it may trap in the pockets of domain II by van der Waals forces and non-covalently interact with TRP-212 and other protein residues; or (ii) it may directly interact with
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Trp-134 and other protein residues in domain I lying on the outer hydrophilic surface of the protein and form a complex. In both cases, quenching of BSA fluorescence is expected to occur either in form of dynamic quenching in first case or static quenching in the second case. To confirm the mechanism of quenching, fluorescence quenching data were analyzed using Stern-Volmer equation Io ¼ 1 þ K sv ½Q ¼ 1 þ K q τ ½Q I
(5)
where, Io and I are steady-state fluorescence intensities in absence and presence of quencher (pDNA) respectively, [Q] is the concentration of quencher, Kq is the quenching rate constant of biomolecule, τo is average life time of biomolecule without quencher and, for BSA its value is 10–8 s (43,44). Ksv is the Stern-Volmer dynamic quenching constant and calculated from the value of slope of regression curve in Io/I Vs [pUC18] plot. From this, the quenching constant Kq can be calculated (Table 2; Fig. S4 and Tables S6 and S7). It was established that the maximum scattering collision quenching constant of various quenchers with the biopolymer is 2.0 × 1010 L mol–1 s–1 (45). Here, in our study we have obtained protein quenching constant Kq (in 1012 range) greater than Kq of standard scattering procedure which demonstrates that quenching is not initiated by dynamic procedure but rather by complex formation (46). Hence, we propose that pDNA binds with BSA-NPs on the hydrophilic surface rather than inside the cavities, and forms intermediate protein–plasmid assembly (which gets transfected into E. coli DH5α culture). To study the conformational changes in BSA on interaction with pDNA, UV-Vis absorption spectroscopy at 7.4 pH was
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Transfecting pDNA to E. coli
Wavelength (nm)
Figure 2. Emission spectra of (a) pUC18 and (b) pUC18egfp on BSA NPs with increase in concentration. (c) for pBBR1MCS-2, see Fig. S3 of SI.
Table 2. Stern-Volmer parameters for plasmidloaded BSA-NPs Plasmid pUC18 pUC18egfp pBBR1MCS-2
Ksv (avg) L/mol 3.8 × 10–2 6.5 × 10–2 7.64 × 10–2
carried out (Fig. 3). As can be seen from the Fig. 3, that blank albumin NPs strongly absorb at 278 nm due to two overlapping S0 to S1 electronic transitions 1La and 1Lb of tryptophan residue while only pUC18 weakly around 255 nm. BSA-NPs loaded with plasmid also absorb at 278 nm with decrease in intensity without altering λmax. This observation confirms the integrity of protein structure in presence of pDNA. Hence, optical studies support the argument that concentration and size of the pDNA loaded on the BSA-NPs govern the quenching of fluorescence. Apparently, with increase in these parameters quenching also increases. FT-IR spectroscopy is one of the best tools for gaining insight into the mode of binding of BSA-NPs with pDNA. The absorption peak analysis also yields information about protein secondary structure and its various conformations. The specific locations
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Wavelength (nm) Figure 3. UV-Vis spectra of (a) pUC18 (b) pUC18egfp. (c) For pBBR1MCS-2 see Fig. S5 of SI.
of interaction and its effect on flexibility of the DNA helical strands can also be predicted. Fig. 4 shows the transmission spectra of blank BSA-NPs and loaded with different plasmids. The peak near 1650 cm–1 is the amide I band. It results from the C = O stretching vibrations of the peptide bond. Similarly, near 1534 cm–1 (N–H bending vibration/C–N stretching vibration) and 1243 cm–1 (C–N stretching vibration/N–H bending vibration) are called the amide II and amide III bands, respectively. The peak near 3400 cm–1 is thought to be N-H bending vibration (also responsible for H-bonding) and at 1400 cm–1 results from protein side-chain COO– (47). Plasmid DNA, in its β-conformation, shows PO2 asymmetric and symmetric stretching at 1225 and 1086 cm–1 respectively. From the optical studies it may be assumed that plasmid interacts with –NH2 groups present on protein surface through its backbone (48). Due to this, the intensities and positions of the corresponding peaks should be changed and shifted. As, in this case, the phosphate asymmetric stretch and N–H bending regions are closely overlapped it is difficult to predict about site of interaction precisely from these regions. However, the intermolecular Hbonding region, 3000–3500 cm–1 can be helpful for the purpose. The broad peak at 3317 cm–1 as seen in Fig. 4 suggests Hbonding among various amino acid residues in bulk BSA protein. This peak gets blue shifted (3446 cm–1) and shallower (V-shape) (49) indicating more compactness and decrease in distance of H-
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Figure 4. Interaction of protein NPs with pDNA. FTIR spectra of (a) Unloaded BSA NPs (b) BSA bulk crystals (c) pUC18 (d) pUC18egfp and (e) pBBR1MCS-2 loaded BSA NPs.
bonded residues on transiting to nano phase. On loading pDNA to protein, the shape of the peak remained the same with red shifting. This observation further supports our argument that pDNA makes a stable complex with protein (50): the shape of the peak remains the same (otherwise it would have changed if pDNA occupied the place in the protein cavities). The plasmid release profile was monitored with respect to time by using a spectrophotometer and comparing the observation with a pUC18 standard curve. It can be observed from Fig. 5 (a–c) that percentage release of pDNA increases with respect to time. It takes a maximum of 240 min to release pDNA from
protein NPs. In the case of pBBR1MCS-2 (5.1 kb) slow release is observed for the initial 100 min followed by a quick release up to another 80 min, while in case of pUC18 (2.9 kb), there is continuous release up to 210 min when it becomes steady. In the case of pUC18egfp steady and stepwise release profile was observed up to 200 min. Hence, it is inferred that percentage release of pDNA directly depends on the size of pDNA. Larger pDNA makes the protein–pDNA complex more labile while smaller pDNA makes a more stable complex. So when allowed to be released, pBBR1MCS-2 gets released with uneven bursts while pUC18 reveals a steady release profile (51).
Figure 5. Release profile of (a) pUC18 and (b) pUC18egfp (c) pBBR1MCS-2 loaded BSA nanoparticles.
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Transfecting pDNA to E. coli For the confirmation of transfection, pUC18 was tagged with enhanced green fluorescence protein, egfp (pUC18egfp) and transfected in E. coli DH5α strain using BSA-NPs as vehicle. It can be observed from Fig. 6 that E. coli harboring pUC18egfp shows fluorescence property under confocal microscope while other plasmids (pUC18 and pBBR1MCS- 2) were unable to show fluorescence due to absence of egfp. This observation endorses that the transfection of E. coli with pDNA can be effectively carried out by using protein NPs as vehicle. It is logical to think that successful transfection should result in expression of specific antibiotic resistance in bacteria. For the purpose, the transfected E. coli DH5α strain were grown in ampicillin comprising plate. It can be perceived from Fig. 7 that the pDNA transfected colonies show growth in antibiotic plates to which they were susceptible formerly. Additionally, to prove the integrity of plasmids after transfection they were recovered from the transfected cells and analyzed. These results also provide evidence for retention of integrity of pDNA after transfection (Fig. 7b). This proves that BSA-NPs are promising candidate for successful gene delivery. The observed transfection efficiency of BSA-NPs could be explained on the basis of “facial amphiphilicity” concept
Figure 7. (a) Growth of antibiotic sensitive E. coli DH5α in diluted LB broth containing ampicillin after 1 day incubation with Plasmid loaded nanoparticles. (b) Plasmid pDNA isolated from transfected E. Coli DH5α. Lane 1- pUC18egfp; Lane 2- pBBR1MCS-2; Lane 3- pUC18 and Lane 4- negative control.
(52,53). BSA protein with outer hydrophilic surface and inner hydrophobic pockets (54), acts as facial amphiphile and can therefore easily transverse the cell membrane.
Figure 6. Confocal microscope images of DH5α transfected with pDNA. A-lane fluorescent filed and B-lane merges of fluorescence and bright field.
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Conclusion BSA-NPs were successfully synthesized by coacervation method. These BSA-NPs were employed as gene-delivery vehicles to load pDNA such as pUC18 carrying ampicillin or kanamycin resistance genes. This protein-conjugated pDNA was successfully and integrally transfected into E. coli DH5α bacterial cells. From this study it can be concluded that: (i) pDNA makes stable complex with protein on its surface by intermolecular H-bonding rather than a simple entrapment inside cavities; (ii) the greater the size of the plasmid, the faster is its release profile. Hence BSA-like natural polypeptides can serve as non-toxic, biodegradable and safe vector to load variety of genes of useful products like enzymes, proteins, peptides, hormones etc into bacterial cells. Such a carrier can also be used for delivering genes into eukaryotic cells and can be a potential candidate for use in gene therapy. This work may further be extended in the fields such as bio-augmentation, DNA profiling, development of biosensors. This study may open a new direction in research towards the development of natural polypeptide-based universal delivery vectors.
13.
14. 15. 16.
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Acknowledgements
22.
Authors are thankful to the Department of Biotechnology, Government of India for providing confocal microscopic images under DBT-MSUB-Interdisciplinary Life Science Programme for Advanced Research and Education.
23. 24. 25.
Author contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.
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Transfecting pDNA to E. coli 38. Engebrecht J, Brent R, Mustak AK. Escherichia coli, plasmids, and bacteriophages: vectors derived from plasmids. Curr Protoc Mol Biol 1991;15:1.6.1–1.6.10. 39. Sambrook J, Russell DW. Molecular cloning: A laboratory manual, 3rd edition. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press, 2001. 40. Gelamo EL, Silva CHTP, Imasato H, Tabak M. Interaction of bovine (BSA) and human (HSA) serum albumin with ionic surfactants: spectroscopy and modeling. Biochim Biophys Acta 2002;1594: 84–99. 41. Lakowicz JR. Principles of fluorescence spectroscopy, 3rd edition. New York: Springer, 2006. 42. Peter T. Serum albumin. Adv Protein Chem 1985;37:161–245. 43. Lackowicz JR, Weber G. Quenching of fluorescence by oxygen: a probe for structural fluctuation in macromolecule. Biochemistry 1973;12:4161–70. 44. Eftink MR. Fluorescence quenching reactions: Probing biological macro-molecular structures. In: Dewey, T.G., editor. Biophysical and biochemical aspects of fluorescence spectroscopy. New York: Plenum Press, 1991. 45. Ware WR. Oxygen quenching of fluorescence in solution: an experimental study of the diffusion process. J Phys Chem 1962; 66:455–58. 46. Anbazhagan V, Renganathan R. Study on the binding of 2,3diazabicyclo[2.2.2]oct-2-ene with bovine serum albumin by fluorescence spectroscopy. J Lumin 2008;128:1454–70. 47. FTIR spectroscopy of protein structure is found at http://www.shimadzu. com/an/industry/pharmaceuticallifescience/proteome0205005.htm.
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48. Tajmir-Riahi HA, N’soukpoé-Kossi CN, Joly D. Structural analysis of protein–DNA and protein–RNA interactions by FTIR, UVvisible and CD spectroscopic methods. Spectroscopy 2009; 23: 81–101. 49. Clayden J, Greeves N, Warren S, Wothers P. Organic chemistry. New York: Oxford University Press, 2001. 50. Dev SB, Walters L. Fourier transform infrared spectroscopy of the characterization of a model peptide–DNA interaction. Biopolymers 1990;29:289–299. 51. Bacalocostantis I, Mane VP, Kang MS, Goodley AS, Muro S, Kofinas P. Effect of thiol pendant conjugates on plasmid DNA binding, release, and stability of polymeric delivery vectors. Biomacromolecules 2012;13:1331–39. 52. Cheng Y, Douglas MH, Gottlieb CR, Kahne D. Facial amphiphiles. J Am Chem Soc 1992;114:7319–20. 53. Mellet CO, Benito JM, Garcia Fernandez JM. Preorganized, macromolecular, gene-delivery systems. Chem Eur J 2010;16: 6728–6742. 54. Hollfelder F, Kirby AJ, Tawfik DS. Off-the-shelf proteins that rival tailor-made antibodies as catalysts. Nature 1996;383:60–3.
SUPPORTING INFORMATION Additional supporting information may be found in the online version of this article at the publisher’s web site.
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Revised: 13 August 2014,
Accepted: 4 September 2014
Published online in Wiley Online Library
(wileyonlinelibrary.com) DOI 10.1002/bio.2789
Transfecting pDNA to E. coli DH5α using bovine serum albumin nanoparticles as a delivery vehicle Jitendra Wagh,b Kuldeep J. Patel,c Parth Soni,c Krutika Desai,c Pratik Upadhyayd and Hemant P. Sonia* ABSTRACT: We describe the formulation of bovine serum albumin nanoparticles (BSA-NPs) by the coacervation method using surfactants. Plasmids (pUC18, pUC18egfp and pBBR1MCS-2) isolated from E. coli were incorporated into the BSA matrix by incubating in albumin solution prior to formulation of NPs. Plasmid incorporation was calculated by % yield, entrapment efficiency, DNA loading capacity and release of entrapped DNA by comparing with blank NPs. BSA-DNA binding studies were carried out by using fluorescence spectroscopy and Fourier Transform Infra Red Spectroscopy (FT-IR). The surface charge distribution of the NPs loaded with plasmid was calculated using zeta potential. The photoluminescence of BSA-NPs was quenched when loaded with pDNA, confirming the interaction of DNA with BSA. Altogether, these results provide evidences for the excellent DNA carrying efficiency of BSA-NPs without loss of plasmid’s integrity. The NPs were used to transfect E. coli DH5α strain lacking ampicillin resistance. They, however, showed ampicillin resistance subsequent to transfection with plasmid encoding ampicillin resistance gene. Effect of transfection was confirmed by confocal microscopy and by the isolation of the plasmid by agarose gel electrophoresis from the transfected bacterial culture. This study clearly demonstrates the efficacy of BSA-NPs as delivery vehicle for pDNA transfection. Copyright © 2014 John Wiley & Sons, Ltd. Additional supporting information may be found in the online version of this article at the publisher’s web site. Keywords: BSA nanoparticles; Gene delivery; Enhanced green fluorescence protein
Introduction Transfection of foreign plasmid DNA or RNA into host cell nucleus to modify, change, or silence expression of gene is a challenging task. Availability of strategy in this context can ensure the production of specific protein that can help to mitigate conditions like Parkinson’s disease (1) and cystic fibrosis (2) or even provide relief to painful chemotherapy treatments in cancer (3). The cellular nuclease enzymes constitute the major obstacle in this task as they can degrade naked plasmids in the cytoplasm before entry into the nucleus leading to low transfection efficiency (4–6). Various strategies have been devised to overcome this problem like trapping such pDNA in a specialized carrier of both natural and synthetic origin (7). Such a carrier should not only bind with pDNA but also with receptors on the cell membrane for successful entry into cells. After entering cytoplasm, it should neither interact with other organelles nor interfere with any biological process (8). Ideally, it should be degraded once it has entered the cytoplasm and release the cargo without causing any harm to the cell. There are carriers that can directly traverse the cell membrane and enter the cytoplasm and viruses are the naturally evolved machinery ideal for such a process (9,10). However, there are serious drawbacks with such systems like with immunogenicity (11,12), oncogenicity (13) and recombination efficiency (14), which restrict the successful application of them. Many chemical agents such as cationic lipids like 2,3-di-oleoyloxytrimethyl ammonium propane (15), fullerenes and their derivatives (16,17), biodegradable polymers and dendrimer based materials e.g. PLL-b-PEG (18–20), PAGA (21), PAMAM type dendrimers etc.; carbohydrate based polymeric cages e.g. β-cyclodextrins (22,23), chitosan (24,25) etc. have been
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considered as nonviral gene-delivery vectors. But they also have their own drawbacks, e.g. high concentrations of fullerene-based materials can induce inflammation promoting the development of cancer (26). Recent reviews have focused on different types of gene-delivery systems, barriers encountered during nonviral gene delivery and the techniques to overcome them (27,28). Considering these, there is a need for development of efficient and specific delivery vehicles either natural or synthetic. Protein, being a macromolecule, can very well satisfy all such requirements to act as a gene-delivery vector (29). They are capable of selfassembly into various shapes and morphologies with well defined
* Correspondence to: Dr. Hemant P Soni. Department of Chemistry, Faculty of Science, The Maharaja Sayajirao University of Baroda, Vadodara-390 002, Gujarat, India. E-mail: [email protected] a
Department of Chemistry, Faculty of Science, The Maharaja Sayajirao University of Baroda, Vadodara-390 002, Gujarat, India
b
Department of Biochemistry, Faculty of Science, The Maharaja Sayajirao University of Baroda, Vadodara-390 002, Gujarat, India
c
Department of Biotechnology, Ashok & Rita Patel Institute of Integrated Study & Research in Biotechnology and Allied Sciences (ARIBAS), Sardar Patel University, V V Nagar, Gujarat, India
d
LJ Institute of Pharmacy, Sarkhej, Ahmedabad, Gujarat, India Abbreviations: AAV, adeno-associated virus; BSA, bovine serum albumin; BSA-NPs, bovine serum albumin nanoparticles; DLS, Dynamic Light Scattering; ESGT, European Society of Gene Therapy; FTIR, Fourier transform infra red; LAFU, laminar air flow unit; LB, Luria Broth.
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J. Wagh et al. pockets. For example, ferritin, a natural protein, has a cavity of 12 nm outer diameter and 8 nm inner diameter (30), pDNA can easily be accommodated intact in such a cavity. They can also be degraded naturally once their function is finished without causing any side effects, even more, their disintegration can be controlled and made more site specific. Hence, proteins have the potential to function as safer, non-toxic, stable and biodegradable genedelivery vectors (31). The purpose of this study was to develop a novel transfection technique by using BSA-NPs. At the initial stage, to demonstrate the efficiency of the developed protein nanoparticles as vehicle, we selected bacterial cells as host to deliver pDNA. However, this strategy may be further extended to eukaryotic cells as well. It has been observed that bacteria can take up protein nanoparticles as a part of their nutrient requirements (32,33). This can therefore, be exploited to transfer biomolecules like pDNA into a bacterium. The entrapment of pDNA in such protein nanoparticles may be an effective means to transfect the gene of interest into a host bacterium. Three different plasmids pUC18 (34), pUC18egfp (35) and pBBR1MCS2 (36) encoding kanamycin and ampicillin resistance genes, were isolated from E. coli culture. These plasmids were eluted, purified and utilized for loading on BSA-NPs. Such gene loaded BSA-NPs were then transfected into E. coli DH5α. As an evidence of successful transfection, bacteria under study were tested for the expected resistance against the specific antibiotics (37). Enhanced green fluorescence protein tag was attached with pUC18 as marker and confocal microscopy of the pUC18egfp transfected E. coli DH5α strain was carried out to confirm successful transfection. In this work, we have presented direct evidence of cell transfection by plasmid-loaded BSA nanoparticles.
Formulation of nanoparticles by coacervation method BSA (2% w/v) solution was prepared in water at 5.5 pH and added to pDNA (Table S2) and incubated for 1 h at room temperature. Then, the aqueous phase was dissolved in chilled ethanol drop wise (2:1, ethanol: water) at a rate of 1 ml/min. The coaservates, so formed were hardened with 25% glutaraldehyde solution (1.56 μg/mg) and further incubated for 2 h. The resulting nanoparticles were purified by centrifugation at 15,000 rpm for 10 min at 4 °C. The supernatant obtained was transferred in another microcentrifuge tube to estimate the amount of protein and unloaded DNA. The resultant pellet of purified nanoparticles was dried and stored at –20 °C. Transfection of E. coli DH5α with plasmid-loaded BSA nanoparticles For the uptake of nanoparticles by E. coli DH5α, pure culture (50 μL) was inoculated in LB (1 g/100 ml instead of 2 g/100 ml) containing filtered NPs solution (200 μL), under LAFU. The culture was allowed to grow over night at 37 °C with constant shaking (Table S3 and S4).
Experimental Materials Potassium acetate, glacial acetic acid, phenol, chloroform, sodium dodecyl sulphate (SDS), 25% glutaraldehyde were purchased from Merck. The restriction enzyme EcoRI, ampicillin, kanamycin, 10× assay buffers and other required chemicals were purchased from the Genei Chemicals, India. Plasmids, pBBR1MCS-2 (a plasmid originally derived from the pBBR1 plasmid of Bordetella bronchiseptica) express kanamycin resistance while, pUC18 and pUC18egfp show ampicillin resistance, which are high copy number plasmids (Table S1). Characterization of pDNA The bacterial cells were harvested to obtain the culture for isolation of plasmid DNA. Single colonies from pure culture were collected in a wire loop and inoculated in 100 ml autoclaved LB broth containing required antibiotic (5 μg/ml). The sample was incubated at 37 °C with 200 rpm overnight for uniform aeration and growth. Plasmid was isolated from bacteria by miniprep method (38). The isolated plasmid was detected by agarose gel electrophoresis (39). The observed bands of plasmid DNA were separated using a sharp sterilized blade, and added to a clean and dry microcentrifuge tube containing fixed weight of gel. In accordance with the weight of gel, three volumes of QG buffer was added. The tube was incubated at 50 °C for 10 min to dissolve the gel. To this, 1 volume of isopropanol was added and mixed thoroughly. The sample was now applied on to the column and centrifuged for 1 min, at 4 °C at 13,000 rpm. To this,
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750 μL of PE buffer was added and the column was allowed to stand for 3 min, followed by centrifugation for 1 min. The flow through was discarded and the column was placed in a clean microcentrifuge tube and allowed to air dry for 5 min. The eluted and purified pDNA was further used for restriction digestion using EcoRI, whose specific restriction sites are present in the plasmids. This helped in size determination of the isolated plasmids. For restriction digestion, the following cocktail was prepared for each sample of DNA consisting of template 50 μg, assay buffer (10×) 2 μL, BSA (10×) 2 μL and EcoRI (10 U/μL). In order to make up the total volume to 20 μL, autoclaved distilled water was added. These mixtures were incubated for 4 h at 37 °C and there after samples were run on 1% agarose gel.
Screening of transfectants To screen successful transfectants, the culture was aspirated (50 μL) and added to LB containing antibiotics for the plasmids used in study. This was allowed to grow over night at 37 °C. Successful growth of bacteria confirmed the uptake of plasmid from NPs with ampicillin resistance. This was further established by performing plasmid isolation using the culture which initially lacked any plasmid. For additional confirmation of successful transfection, bacterial cells were washed with normal saline for egfp expression present in pUC18egfp and then confocal microscopy (Carl Zeiss Microscopy) was carried out at excitation wavelength 485 and emission at 510 nm.
Characterization and evaluation Evaluation of pDNA The concentration and purity of plasmid DNA was measured in a UV spectrophotometer. For the purpose, pDNA solution (50 μL) was mixed in distilled water (2950 μL) and the absorption spectra were measured at 260 and 280 nm. The concentration of pDNA was expressed as ng/μL. The purity of DNA was assessed by 260/280 nm absorption ratio (Table S5). The pDNA was subjected to restriction digestion using EcoRI enzyme and agarose electrophoresis was carried out to determine the size
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Transfecting pDNA to E. coli of the eluted pDNA. The linear DNA fragments detected after digestion were separated on a 1% agarose gel along with the reference ladder.
pDNA added and the amount present in the supernatants obtained after the purification by spectrophotometer at 260 nm. DNA loading capacity was calculated as,
Evaluation of nanoparticles
% DNA loading capacity ¼
The mean size of formulated NPs was determined by Dynamic Light Scattering (DLS) technique. A BIC 90 plus (Brookhaven) equipped with 35.0 mW solid-state laser operating at 660 nm and an avalanche photodiode detector was used for the measurement of particle size distribution by DLS and surface charges were measured in term of zeta potential by a Zeta analyzer system, Nano Series (Malvern Instruments). The surface morphology and size of NPs formulations were also studied by Transmission Electron Microscopy operating at 200 kV (Philips CM200). The amount of protein transformed in NPs was determined by standard BCA protein assay. NPs obtained after centrifugation were digested in basic medium for 2 h at RT. The absorbance of the resulting solutions was measured at 562 nm. The% yield of NPs was calculated as, % yield ¼
Total weight of NPs 100 total weight of pDNA and BSA
(1)
%Entrapment efficiency ¼
Amount of DNA in NPsðμgÞ100 Yield of NPsðmgÞ (2)
Amount of pDNA present in NPs Amount of pDNA used in formulation
(3) Plasmid-loaded BSA-NPs were suspended in phosphate buffer saline (PBS, 300 μL) at pH 7.4 as release media in microcentrifuge tubes placed in shaking water bath at 37 °C at 60 rpm. At predetermined intervals, 50 μL samples were harvested and replaced with 50 μL fresh media. For the determination of amount of DNA released, the tubes were centrifuged and supernatant obtained was analyzed by using spectrophotometer at 260 nm. % Release of pDNA ¼
Amount of pDNA released Amount of pDNA present in NPs (4)
BSA-DNA binding study
Results and discussion
The interaction between BSA and pDNA in aqueous solution was investigated using fluorescence spectroscopy by means of Jasco FP-6300 spectrofluorometer using Xenon lamp as the excitation source. UV-Vis absorption spectra were recorded on a Perkin Elmer Lamda 35 uv/vis spectrometer. The binding constant Ksv was determined using Stern-Volmer equation (40). The required, λmax of BSA was measured by UV spectroscopy. This λmax value, in turn, was used to excite loaded and unloaded BSA nanoparticles for emission spectrum in fluorescence spectroscopy, by keeping the concentration of BSA constant, and simultaneously increasing the amount of pDNA. Loading of pDNA on BSA nanoparticles and interaction of pDNA with BSA nanoparticles were also studied. For the purpose, the absorption spectra of pure BSA, pure pDNA, BSADNA solution, blank nanoparticles (only BSA) and pDNA loaded nanoparticles, were obtained by using a UV spectrophotometer. Samples with appropriate concentration were scanned at 200–400 nm keeping water as blank. Fourier transform infrared (FTIR) spectra of the samples were recorded in the region of 500–4000 cm–1 using Perkin Elmer FTIR Spectrometer (Spectrum RX1).
The plasmids (pUC18, pBBR1MCS2 and pUC18egfp) were isolated from E. coli DH5α and loaded on BSA-NPs for cell transfection. An absorption ratio (260:280 nm) around 1.8 indicates that the isolated plasmids are pure and not contaminated by any protein impurity. Per cent yield, pDNA loading capacity and pDNA loaded on the NPs data suggest that BSA can be an ideal protein to serve as vehicle for pDNA transfection (Table S2,S3 and S4). TEM images show BSA-NPs to be spherical and monodispersed in the size range of 150–170 nm (Fig. 1). It can be observed that on binding pUC18 on BSA-NPs, the size increases in the range of 180–200 nm with almost no change in shape or dispersity. This is also true for other two plasmids as well. The DLS plots for free and bound BSA-NPs (100 vs. 200–300 nm) clearly support the TEM observation (Fig. S2). Zeta potential measurement for free NPs (–53 meV) indicates that the aqueous dispersion of NPs is quite stable. On pDNA binding, the value decreases to lower negative (Table 1) indicating depletion of charge and agglomeration (140–200 nm) of NPs (37). From the zeta potential values, the mechanism of interaction of BSA-NPs with pDNA may be predicted. While in the aqueous medium at acidic pH (5.5), the N and C terminals of the protein are positively and negatively charged respectively, the negative charge gets partially neutralized on pDNA binding due to which a marked depletion of charge occurs in the bound NPs. On comparing Tables 1 and S2 it can be seen that the size of the plasmid also matters for the depletion of charge. As the size increases, more charge on the carrier neutralizes resulting in depletion of charge of the overall protein-plasmid assembly. Fluorescence spectroscopy is an important tool to study the interaction among biomolecules and NPs. It can not only suggest the mechanism of interaction but also provide clues about the micro-environment of fluorophore (BSA in this case) from which fluorescence originates. It can also provide information
Evaluation of plasmid DNA loaded nanoparticles The absorbance of BSA solution with different concentrations in basic medium was measured by UV spectrophotometer at 562 nm using 0.1 M NaOH solution as blank. From the data, a calibration curve of concentration versus absorbance was plotted (Fig.S1). Further, standard calibration curve was obtained for eluted pDNA solution in water at 260 nm. Amount of pDNA released from nanoparticles can be estimated on comparing the curves obtained by UV spectrometry with the standard curve. The amount of pDNA loaded on BSA nanoparticles was calculated as the difference between the total amount of the initial
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Figure 1. TEM images of (a) unloaded (b) pUC18 (c) pBBR1MCS-2 and (d) pUC18egfp loaded BSA nanoparticles.
Table 1. Zeta potential of BSA nanoparticles loaded with different plasmids Nanoparticles Loaded with size of the Zeta potential plasmid plasmid in kb (mV) BSA BSA BSA BSA
– pUC18 pUC18egfp pBBR1MCS-2
– 2.9 3.5 5.1
–53.2 –41.6 –9.04. –10.4
about conformational changes taking place when protein interacts with other biomolecules (41). Tryptophan (Trp), tyrosine (Tyr) and phenylalanine (Phe) residues are mainly responsible for fluorescence in BSA. While one tryptophan residue (Trp134) is located in the first domain the other (Trp-212) is located in the second domain of BSA protein. Trp-212 is situated inside the hydrophobic binding pocket while TRP-134 is on the outer hydrophilic surface of the BSA (42). Fig. 2 shows the fluorescence emission spectra of BSA in presence of different concentrations of pUC18, pUC18egfp and pBBR1MCS-2 following an excitation at 280 nm. It can be seen from Fig. 2 that BSA fluoresces strongly at 675 nm when excited at 280 nm at pH 7.4 and its intensity gradually decreases with increasing concentration of pDNA. Further, the blue shifting in the λmax value occurs with increase in concentration. When pDNA comes in the vicinity of the protein, there are two possibilities: (i) it may trap in the pockets of domain II by van der Waals forces and non-covalently interact with TRP-212 and other protein residues; or (ii) it may directly interact with
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Trp-134 and other protein residues in domain I lying on the outer hydrophilic surface of the protein and form a complex. In both cases, quenching of BSA fluorescence is expected to occur either in form of dynamic quenching in first case or static quenching in the second case. To confirm the mechanism of quenching, fluorescence quenching data were analyzed using Stern-Volmer equation Io ¼ 1 þ K sv ½Q ¼ 1 þ K q τ ½Q I
(5)
where, Io and I are steady-state fluorescence intensities in absence and presence of quencher (pDNA) respectively, [Q] is the concentration of quencher, Kq is the quenching rate constant of biomolecule, τo is average life time of biomolecule without quencher and, for BSA its value is 10–8 s (43,44). Ksv is the Stern-Volmer dynamic quenching constant and calculated from the value of slope of regression curve in Io/I Vs [pUC18] plot. From this, the quenching constant Kq can be calculated (Table 2; Fig. S4 and Tables S6 and S7). It was established that the maximum scattering collision quenching constant of various quenchers with the biopolymer is 2.0 × 1010 L mol–1 s–1 (45). Here, in our study we have obtained protein quenching constant Kq (in 1012 range) greater than Kq of standard scattering procedure which demonstrates that quenching is not initiated by dynamic procedure but rather by complex formation (46). Hence, we propose that pDNA binds with BSA-NPs on the hydrophilic surface rather than inside the cavities, and forms intermediate protein–plasmid assembly (which gets transfected into E. coli DH5α culture). To study the conformational changes in BSA on interaction with pDNA, UV-Vis absorption spectroscopy at 7.4 pH was
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Transfecting pDNA to E. coli
Wavelength (nm)
Figure 2. Emission spectra of (a) pUC18 and (b) pUC18egfp on BSA NPs with increase in concentration. (c) for pBBR1MCS-2, see Fig. S3 of SI.
Table 2. Stern-Volmer parameters for plasmidloaded BSA-NPs Plasmid pUC18 pUC18egfp pBBR1MCS-2
Ksv (avg) L/mol 3.8 × 10–2 6.5 × 10–2 7.64 × 10–2
carried out (Fig. 3). As can be seen from the Fig. 3, that blank albumin NPs strongly absorb at 278 nm due to two overlapping S0 to S1 electronic transitions 1La and 1Lb of tryptophan residue while only pUC18 weakly around 255 nm. BSA-NPs loaded with plasmid also absorb at 278 nm with decrease in intensity without altering λmax. This observation confirms the integrity of protein structure in presence of pDNA. Hence, optical studies support the argument that concentration and size of the pDNA loaded on the BSA-NPs govern the quenching of fluorescence. Apparently, with increase in these parameters quenching also increases. FT-IR spectroscopy is one of the best tools for gaining insight into the mode of binding of BSA-NPs with pDNA. The absorption peak analysis also yields information about protein secondary structure and its various conformations. The specific locations
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Wavelength (nm) Figure 3. UV-Vis spectra of (a) pUC18 (b) pUC18egfp. (c) For pBBR1MCS-2 see Fig. S5 of SI.
of interaction and its effect on flexibility of the DNA helical strands can also be predicted. Fig. 4 shows the transmission spectra of blank BSA-NPs and loaded with different plasmids. The peak near 1650 cm–1 is the amide I band. It results from the C = O stretching vibrations of the peptide bond. Similarly, near 1534 cm–1 (N–H bending vibration/C–N stretching vibration) and 1243 cm–1 (C–N stretching vibration/N–H bending vibration) are called the amide II and amide III bands, respectively. The peak near 3400 cm–1 is thought to be N-H bending vibration (also responsible for H-bonding) and at 1400 cm–1 results from protein side-chain COO– (47). Plasmid DNA, in its β-conformation, shows PO2 asymmetric and symmetric stretching at 1225 and 1086 cm–1 respectively. From the optical studies it may be assumed that plasmid interacts with –NH2 groups present on protein surface through its backbone (48). Due to this, the intensities and positions of the corresponding peaks should be changed and shifted. As, in this case, the phosphate asymmetric stretch and N–H bending regions are closely overlapped it is difficult to predict about site of interaction precisely from these regions. However, the intermolecular Hbonding region, 3000–3500 cm–1 can be helpful for the purpose. The broad peak at 3317 cm–1 as seen in Fig. 4 suggests Hbonding among various amino acid residues in bulk BSA protein. This peak gets blue shifted (3446 cm–1) and shallower (V-shape) (49) indicating more compactness and decrease in distance of H-
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Figure 4. Interaction of protein NPs with pDNA. FTIR spectra of (a) Unloaded BSA NPs (b) BSA bulk crystals (c) pUC18 (d) pUC18egfp and (e) pBBR1MCS-2 loaded BSA NPs.
bonded residues on transiting to nano phase. On loading pDNA to protein, the shape of the peak remained the same with red shifting. This observation further supports our argument that pDNA makes a stable complex with protein (50): the shape of the peak remains the same (otherwise it would have changed if pDNA occupied the place in the protein cavities). The plasmid release profile was monitored with respect to time by using a spectrophotometer and comparing the observation with a pUC18 standard curve. It can be observed from Fig. 5 (a–c) that percentage release of pDNA increases with respect to time. It takes a maximum of 240 min to release pDNA from
protein NPs. In the case of pBBR1MCS-2 (5.1 kb) slow release is observed for the initial 100 min followed by a quick release up to another 80 min, while in case of pUC18 (2.9 kb), there is continuous release up to 210 min when it becomes steady. In the case of pUC18egfp steady and stepwise release profile was observed up to 200 min. Hence, it is inferred that percentage release of pDNA directly depends on the size of pDNA. Larger pDNA makes the protein–pDNA complex more labile while smaller pDNA makes a more stable complex. So when allowed to be released, pBBR1MCS-2 gets released with uneven bursts while pUC18 reveals a steady release profile (51).
Figure 5. Release profile of (a) pUC18 and (b) pUC18egfp (c) pBBR1MCS-2 loaded BSA nanoparticles.
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Transfecting pDNA to E. coli For the confirmation of transfection, pUC18 was tagged with enhanced green fluorescence protein, egfp (pUC18egfp) and transfected in E. coli DH5α strain using BSA-NPs as vehicle. It can be observed from Fig. 6 that E. coli harboring pUC18egfp shows fluorescence property under confocal microscope while other plasmids (pUC18 and pBBR1MCS- 2) were unable to show fluorescence due to absence of egfp. This observation endorses that the transfection of E. coli with pDNA can be effectively carried out by using protein NPs as vehicle. It is logical to think that successful transfection should result in expression of specific antibiotic resistance in bacteria. For the purpose, the transfected E. coli DH5α strain were grown in ampicillin comprising plate. It can be perceived from Fig. 7 that the pDNA transfected colonies show growth in antibiotic plates to which they were susceptible formerly. Additionally, to prove the integrity of plasmids after transfection they were recovered from the transfected cells and analyzed. These results also provide evidence for retention of integrity of pDNA after transfection (Fig. 7b). This proves that BSA-NPs are promising candidate for successful gene delivery. The observed transfection efficiency of BSA-NPs could be explained on the basis of “facial amphiphilicity” concept
Figure 7. (a) Growth of antibiotic sensitive E. coli DH5α in diluted LB broth containing ampicillin after 1 day incubation with Plasmid loaded nanoparticles. (b) Plasmid pDNA isolated from transfected E. Coli DH5α. Lane 1- pUC18egfp; Lane 2- pBBR1MCS-2; Lane 3- pUC18 and Lane 4- negative control.
(52,53). BSA protein with outer hydrophilic surface and inner hydrophobic pockets (54), acts as facial amphiphile and can therefore easily transverse the cell membrane.
Figure 6. Confocal microscope images of DH5α transfected with pDNA. A-lane fluorescent filed and B-lane merges of fluorescence and bright field.
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J. Wagh et al.
Conclusion BSA-NPs were successfully synthesized by coacervation method. These BSA-NPs were employed as gene-delivery vehicles to load pDNA such as pUC18 carrying ampicillin or kanamycin resistance genes. This protein-conjugated pDNA was successfully and integrally transfected into E. coli DH5α bacterial cells. From this study it can be concluded that: (i) pDNA makes stable complex with protein on its surface by intermolecular H-bonding rather than a simple entrapment inside cavities; (ii) the greater the size of the plasmid, the faster is its release profile. Hence BSA-like natural polypeptides can serve as non-toxic, biodegradable and safe vector to load variety of genes of useful products like enzymes, proteins, peptides, hormones etc into bacterial cells. Such a carrier can also be used for delivering genes into eukaryotic cells and can be a potential candidate for use in gene therapy. This work may further be extended in the fields such as bio-augmentation, DNA profiling, development of biosensors. This study may open a new direction in research towards the development of natural polypeptide-based universal delivery vectors.
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Acknowledgements
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Authors are thankful to the Department of Biotechnology, Government of India for providing confocal microscopic images under DBT-MSUB-Interdisciplinary Life Science Programme for Advanced Research and Education.
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Author contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.
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