Solvent mixture used in particle formation

Since 6,13-bis(triisopropylsilylethynyl) pentacene (TIPS pentacene), DNA, and PLGA have very different solubility characteristics, a solvent mixture was needed to solubilize all three components and form well-defined nanoparticles when mixed with the aqueous Pluronic F-127 solution. TIPS pentacene is highly nonpolar and therefore readily soluble in THF while only partially soluble in DMF (Fig. 1a, b). In contrast to TIPS pentacene, PLGA was more soluble in DMF than THF. However, DNA was not soluble in this purely organic mixture and thus a mixed aqueous-organic mixture was sought. Ke et al. [15] used a mixture of 5 vol% TE buffer and 95 vol% DMF and showed that plasmid DNA was stable in this mixture at room temperature. Therefore, to solubilize the DNA, some TE buffer was added to the mixture of DMF and THF but the concentration was kept low to prevent the TIPS pentacene from precipitating. Because the CRISPR plasmid used in this work was about twice the size as that used by Ke et al., 5 vol% was not sufficient to solubilize the DNA and was found experimentally to leave a small visible pellet of DNA that was not loaded into the syringe for particle formation. The final solvent mixture solubilizing TIPS pentacene, DNA, and PLGA consisted of 10 vol% TE buffer, 45 vol% DMF, and 45 vol% THF and was used in the nanoprecipitation process to form nanoparticles.

Fig. 1 TIPS pentacene can be used as a fluorescent marker for cell internalization. a TIPS structure, b normalized absorbance spectrum of TIPS pentacene in THF (v/v), c calibration curve of absorbance vs. TIPS pentacene concentration in THF by volume, d representative imaging flow cytometer pictures selected randomly out of 10,000 BMDM cells treated with either mock or TIPS loaded NPs demonstrates internalization of particles into the cell as indicated by red fluorescence signal Full size image

Evaluation of nanoparticles made with different PLGA end groups show main difference to be in DNA loading

For particles made with the ester-end capped PLGA, the intensity-average hydrodynamic diameters (D i ) were ~ 160 nm while the diameters of particles made with the amine-end capped PLGA were slightly larger, D i ~ 180 nm with no significant change when DNA was encapsulated (Table 1). Because PLGA degrades by hydrolysis, the particles were freeze dried for extended storage which greatly simplified the subsequent cell-based experiments. Since PLGA aggregates irreversibly during freeze-drying, trehalose was used as a cryoprotectant. A mass ratio of NP to trehalose ranging from 1:25 to 1:42—determined by experiments described in Additional file 1—resulted in particles redispersed in DI water that were somewhat aggregated during freeze drying with D i ~ 210–350 nm but still small enough to be useful for cell uptake. The trehalose:NP ratio varied slightly from batch to batch as the exact concentration of the final suspension after the centrifugal wash during particle fabrication varied and was calculated after the freeze-drying process. The zeta potentials of the particles showed little variation, ranging from − 29 to − 35 mV, due most likely to carboxylate groups on the particles formed due to hydrolysis of the PLGA.

Table 1 Size and zeta potential of NPs in DI water made with different end-capped PLGA before and after lyophilization Full size table

Proton NMR analysis can be used to estimate Pluronic F127 versus PLGA mass ratio in nanoparticles

Because Pluronic F127 was added in excess and any material not physisorbed to the surfaces of the nanoparticles during formation was removed during the centrifugation step, it was important to determine the polymer composition of the final nanoparticle product. By determining the mass content of F127 and PLGA in the nanoparticles, the encapsulation efficiency can also be more accurately calculated. Using proton NMR, solutions of the PLGA and F127 in deuterated chloroform (CDCl 3 ) separately were first analyzed.

In the 1H NMR spectrum of Pluronic F127, the methyl protons have a chemical shift around 1 ppm (Fig. 2a). All other peaks from both PEO and PPO blocks are integrated into one peak, between 3.2 and 3.8 ppm, due to the presence of adjacent oxygen atoms. Based on the NMR, the composition of the F127 is PEO 108 -b-PPO 65 -b-PEO 108 which is close to the theoretical values of PEO 100 -b-PPO 65 -b-PEO 100 [16]. Alternatively, if the integrals of the methyl protons on the PPO segment are set to be 100, then there would be 100/3 = 33.3 repeating units of PO. The PEO methylene proton peaks then have an integral of 544.2 – 100 = 444.2, after subtracting the methylene and methine protons from the PO. It means there are 444.2/4 = 111 repeating units of EO, per 33.3 units of PO. The molar ratios of the EO over PO are then:

Fig. 2 Proton NMR of materials making up nanoparticles. Analysis of F127 and PLGA proton NMR spectra show distinct peaks that can be used to determine polymer composition of resulting nanoparticles. 1H NMR spectra of a Pluronic F127 and b ester end capped PLGA c amine end capped PLGA in CDCl 3 Full size image

$$\frac{mole\,of\,EO}{mole\,of\,PO}=\frac{3.33 }{1}$$

Another important value was the percent of the integrals in the overlapped chemical shift region that contributed to the EO mass. The % value becomes important when calculating the PPO ratios in case overlapping occurs when mixed with the PLGA components. The PPO proton wt% can be calculated as follows.

$$\%\text{A}_{3.25-3.8\text{ppm}\text{PPO}}=\frac{444.2}{544.2}=81.2\%$$

The PLGA spectra show two distinctive peaks around 4.5–5.5 ppm (Fig. 2b, c). Specifically, the methine protons (g peak) from the poly(lactide) segment have a chemical shift of ~ 5–5.5 ppm, while the value for the methylene protons (f peak) from the poly(glycolide) segment was 4.5–5 ppm. The molar ratios of the poly(lactide) segment over poly(glycolide) can be determined by comparing the integral of g to the integral of f divided by two, as there are two protons in peak f compared to only one in peak g. The results indicated the molar ratios are ~ 1:1, close to the values provided by the manufacturer.

Because the F127 and the poly(lactide) and poly(glycolide) have distinctive, non-overlapping peaks, the PLGA to F127 ratios in the TIPS loaded NPs could be calculated. For example, using the ester end cap PLGA case (Fig. 3a):

Fig. 3 Using proton NMR to more accurately calculate DNA encapsulation efficiency. Mass ratio of Pluronic F127 and PLGA in the nanoparticles determined by proton NMR allows for more accurate calculations of encapsulation efficiency. 1H NMR of NPs made with Pluronic F-127 and a with ester end capped PLGA and b with amine end capped PLGA Full size image

$$\frac{\text{mole of EO}}{\text{mole of PO}}=\frac{3.3}{1};\quad \frac{\text{mole of GA}}{\text{mole of LA}}=\frac{\frac{253.3}{2}} {\frac{100}{1}}={\frac{1.27}{1}}; \quad \%\text{A}_{3.25-3.8\text{ppm}\text{PPO}}=81.2\%$$

$$\frac{\text{mole \,of \, EO}}{\text{mole \,of \, LA}}=\frac{\frac{96.3*81.2\%}{4}}{100}=\frac{1}{5.11}=\frac{3.33 \text{EO}}{17.0 \text{LA}}$$

The result means that in the mixed systems, per 3.33 mol of EO, there would be 1 mol of PO, 17 mol of LA, and 17 * 1.27 mol of GA. With all these ratios, we can use the MW of the repeat units to find the mass ratio of F127 to PLGA and from there the wt% of each:

$$\frac{{\mathrm{m}}_{\mathrm{F}127}}{{\mathrm{m}}_{\mathrm{P}\mathrm{L}\mathrm{G}\mathrm{A}}}=\frac{3.33\,*\left(44.04\right)+58.06}{17.0\,*\left(1.27\,*\left(58.02\right)+72.04\right)}=\frac{204.7}{2477.3}\to 91.7\mathrm{\%}\, \mathrm{PLGA}/8.3\mathrm{\%}\, \mathrm{F}127$$

This same method can be used for the amine end cap case (Fig. 3b), yielding similar results of 92.4% PLGA. Given the approximate 5% error in the NMR spectra integration, the uncertainty of these calculations is ± 6–7% and therefore both of these compositions are statistically identical and assumed to be ~ 92% PLGA for all future loading calculations.

DNA release profiles differ between NP formulations and pH conditions

From an analysis of the DNA content of the supernatant after centrifugation during the fabrication process, the DNA loading was determined to be 0.7 and 1.6 wt% for the ester and amine endcap PLGA cases, respectively (Table 1). The difference between the two formulations may be due to charge and hydrophobic interactions from the different PLGA end groups. We hypothesized that an amine end group would provide additional electrostatic attraction with the negatively charged DNA which would enhance loading. By contrast, the PLGA with the ester end cap contained a 9-carbon chain as the end group and thus the lower DNA loading with this polymer could be due to a combination of the lack of attractive charge interactions and the increased hydrophobicity of the chain ends which could interact unfavorably with the hydrophilic DNA. Based on the particle sizes and DNA loadings of the samples in Table 1, the estimated number of plasmids per NP ranged from ~ 2 to 5 copies (Additional file 1: Eq. S4).

DNA release measurements were performed at three different pH values (pH 7, 6, 4.5) to mimic the different pH environments that the particles would experience during incubation in the media (pH 7.4) outside the cell, through early (pH 6.8–6.1) and late endocytosis (pH 6.0–4.8), and in lysosomes (pH 4.5) inside the cell [17]. The amine end-capped PLGA case shows a higher release at all three pHs, with the highest release being DNA equivalent to 0.8 wt% loading with respect to PLGA (Additional file 1: Eq. S5) at pH 7 after 3 days (Fig. 4). By contrast, the corresponding release for the ester end-capped case after 3 days at pH 7 was 0.4 wt% with somewhat lower values at pH 4.5 and 6.0. Because the amine end-capped case had over 2× higher overall loading, it is reasonable that it would release more DNA relative to the ester end-capped case.

Fig. 4 In-vitro DNA release profile from nanoparticles. Majority of DNA released within the first 24 h with NPs made from amine endcaps at pH 7 showing the highest level of release. DNA release profile from the particles with respect to time at pH a 7.0, b 6.0, c 4.5. % released = [mass DNA released into solution]/[initial total mass DNA encapsulated] Full size image

From 1–3 days, release at pH 7.0 appeared to be systematically higher by ~ 30–60% than for the pH 4.5 case. However, it is possible that pH may affect the concentration of DNA detected by the PicoGreen assay. The first step to depurination and β-elimination during dsDNA degradation in aqueous media is catalyzed by acidic conditions [18]. Evans et al. [18] showed in their accelerated stability studies that even at a pH 6, significant difference in degradation could be seen for supercoiled plasmid DNA when compared to pH 7. The formation of acid groups due to hydrolysis is probably why lower DNA release was measured for the lower pH cases. Hydrolysis of PLGA is catalyzed by acidic conditions and, as the PLGA breaks down to form more acid groups, the local pH inside the core decreases [19]. This positive feedback loop accelerates the further breakdown of PLGA [20]. When the pH of the surrounding media was already lower as in the pH 4.5 and 6.0 cases, the acid-catalyzed hydrolysis happens more rapidly and thus could have degraded more DNA than in the more neutral pH 7 case. If the DNA is exposed to these highly acidic conditions for a long period of time, it could degrade quickly and fall below the detection limit of the assay. Balmert et al. [19] estimated the intraparticle pH of ester endcapped PLGA microparticles (MW = 15 kDa) as ~ 3–4 within 1–3 days in neutral pH media conditions. This may account for the relatively rapid release of the DNA. The particles are rapidly being hydrolyzed which, in turn, forms more acid groups that further catalyze hydrolysis, leading to formation of pores that lead to faster diffusion of DNA out of the particle into the aqueous media. However, at lower pH values, especially at pH 4.5, the acidic environment in the nanoparticles may lead to early DNA degradation, thus lowering the apparent release levels. This is supported by the decline in DNA release for the pH 4.5 case after 3 days.

The apparently anomalous points in the release profile at the t = 0 time point at all three pH values occurred when the buffer was initially added to the NPs, followed by immediate centrifugation to obtain the supernatant for the PicoGreen assay. The DNA content at this time point was found to be higher than the subsequent 1-h time point for all six of the DNA-containing conditions tested. We believe this was due to surface-bound or partially encapsulated DNA that may have been partly degraded during processing due to its exposure to the environment. Because that DNA was close to the surface, it was quickly released. Although all processing steps were done carefully to minimize degradation, between mixing, centrifugation, freeze drying, and reconstitution in buffers, it is possible that some of the surface-bound DNA had degraded. We hypothesize that, at t = 0, this surface-bound DNA was released rapidly and detected by the assay. If this DNA had already been partially degraded to form relatively short linear DNA fragments due to the effects of the handling steps, it may have degraded faster once in the media with its chain length eventually falling below the detection limit of PicoGreen (< 200 bp) as specified by the manufacturer. Other sources have shown experimentally that PicoGreen could accurately detect DNA chains as short as 150 bp [21, 22]. Regardless of the cutoff length for detection, the hypothesis of partially degraded DNA chains on the surface undergoing rapid burst release and degradation to lengths below detection by PicoGreen still applies.

This degraded DNA can also show up as a stronger signal for the same amount of DNA than when in plasmid form given the nature of the PicoGreen assay. The assay involves intercalation of the reagents into the DNA and therefore will not have access to the entire chain when the plasmid is supercoiled. Holden et al. [23] reported that, for their plasmid, the PicoGreen assay showed the supercoiled plasmid to be 60% the mass of the same plasmid that had been linearized. The discrepancy between the supercoiled and linearized forms will depend on the sequence and conformation of the plasmid but, in all cases, the supercoiled case may show a lower signal due to inaccessibility of parts of the chain. The DNA concentration for the stock solution was measured by UV absorption using a NanoDrop 2000 (ThermoFisher) which is thought to be more accurate than the PicoGreen assay for plasmids. DNA concentration measurements made with the NanoDrop 2000 were used to concentrate the stock for a targeted 2 wt% DNA loading with respect to mass of PLGA. Under the assumption that the added DNA was enough for exactly 2 wt% DNA loading, the unincorporated DNA and encapsulated DNA should add up to that total mass added. However, given the lowered detection by PicoGreen, the mass of unincorporated DNA as measured from the supernatant using PicoGreen would be an underestimate. Similarly, the DNA released over the 5 days was ~ 50% of what was loaded. These measurements are also underestimates and could be a main factor in accounting for the missing mass in the mass balance.

More important than the actual estimated loading is the DNA released as shown by the release study. The amount of measured DNA released for the amine case after 5 days at pH 7.0 was equivalent to a DNA loading of 0.8 wt% with respect to PLGA or approximately half of the total 1.6 wt% loading. This corresponds to ~ 2–3 plasmid copies released per NP and is a rough underestimate as mentioned above. An underestimated plasmid release is better than an overestimate in this application because the chances of successful delivery of the plasmid to the nucleus for transcription increases with the number of plasmid copies released. Therefore, the particles may be more effective for the apparent DNA added. To test this, cell studies were conducted to investigate the expression of the Cas9 protein and to explore any changes to the mouse DNA after NP treatments.

Bacterial S. pyogenes Cas9 protein is successfully translated inside murine macrophages

To further test the successful encapsulation of CRISPR plasmid into the amine end-capped PLGA nanoparticles, we next wanted to determine whether the plasmid remained functional, defined by its ability to transcribe and translate S. pyogenes Cas9 protein. To do so, we harvested wild type mouse bone marrow derived macrophages (BMDMs), replated at a density of 500,000 cells/mL, and challenged the macrophages with either blank nanoparticles (100 µg/mL), CRISPR plasmid-loaded nanoparticles (100 µg/mL), CRISPR plasmid with Lipofectamine 3000 transfection (2 µg/mL DNA), CRISPR plasmid only (2 µg/mL), or PBS for 24 h. The total remaining cells were removed from the plates, lysed, and Western blot was performed for Cas9 using a S. pyogenes specific monoclonal antibody (Fig. 5). BMDM’s will readily phagocytose plasmids, such as the CRISPR–Cas9 plasmid used here, and Lipofectamine is a common method for plasmid transfection. Thus, we anticipate that both of these methods will successfully introduce the functional CRISPR plasmid into our BMDMs under our experimental conditions. However, neither of these methods are appropriate for in vivo use where Lipofectamine has well known toxicity issues and is not biocompatible. Likewise, the naked plasmid lacks stability and is rapidly degraded once administered in vivo. The goal of these studies is to compare the functionality of the plasmid following nanoparticle delivery against these more typical approaches.

Fig. 5 Bacterial S. pyogenes Cas9 protein is successfully translated inside murine macrophages. After 24 h incubation with CRISPR+ Lipofectamine (lanes 1 and 2), CRISPR plasmid only (lanes 3 and 4), PBS only (lane 5), CRISPR loaded nanoparticle (lane 6), and blank nanoparticle (lane 7), S. pyogenes Cas9 protein was detectable by Western Blot Full size image

The nanoparticle concentrations were chosen to keep the DNA concentration constant between the samples under the assumption of 2 wt% targeted DNA loading. However, the measured loading was 1.6 wt% with respect to the PLGA, and with the presence of F127 included in the total NP mass, the nominal DNA concentration of the plasmid NP case was approximately 1.5 µg/ml. In order to control for suboptimal nanoparticle delivery of CRISPR plasmids, we used Lipofectamine 3000 (Invitrogen) in order to transfect approximately the same total DNA that was encapsulated in the particles. Due to the phagocytic nature of the BMDM primary cells that we used for this study, we also treated the cells with the free plasmid DNA. Cas9 was detectable in the cells transfected with Lipofectamine (lanes 1 and 2) as well as the cells treated with CRISPR plasmid only (lanes 3 and 4) and CRISPR-loaded nanoparticle (lane 6), while the cells treated with blank nanoparticle (lane 7) and PBS only (lane 5) were not (Fig. 5). Qualitatively, the band intensities between all three CRISPR-containing samples were comparable. Again, given the phagocytic nature of these cells, the BMDMs internalized the plasmid-only control with no additional carrier or transfection needed. From release studies shown earlier, we showed that most of the plasmid was released from the particles within the first 24 h in suspension, and more specifically within the first 8 h. From imaging cytometry, we counted 10,000 cells per experimental condition and found 95% of the macrophages treated with nanoparticles exhibited red fluorescence from the TIPS pentacene indicating internalization after 24 h (Fig. 1d). McDaniel et al. showed similar statistics using TIPS pentacene loaded poly(lactic acid)-based nanoparticles. That study also showed that within the first 2 h ~ 30% of cells showed particle uptake increasing to ~ 40% at 4 h but not reaching the 90+% until after 8 h of incubation [24]. Assuming similar DNA release kinetics in cell culture media, and similar particle uptake behaviors with these PLGA particles, it is difficult to discern whether the entire nanoparticle was internalized by the macrophages before releasing the plasmid into the cytosol as intended, the plasmid in the particles were released outside the cell and the free plasmids phagocytosed, or a combination of the two. We hypothesize this will become clearer in future in vivo studies. Cohen et al. [10] found that nanoparticles performed better than liposomes for in vivo delivery of plasmid DNA for gene editing applications, although it did not do as well in in vitro cell studies. Even though we cannot see a clear advantage in using transfecting agents from this particular study, what this result does show is that the encapsulated high molecular weight plasmids in the nanoparticles were intact enough to express the Cas9 protein and can therefore be considered functional. In the current set of studies, we cloned a test gRNA targeting the Lps-d allele in the mouse Tlr4 gene into our pX330 CRISPR plasmid [25]. Future studies will include functional and validated CRISPR gRNAs that target a range of murine genes of interest both in vitro and in vivo.