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JoVE Journal Bioengineering

Bioengineering

Formulating and Characterizing Lipid Nanoparticles for Gene Delivery using a Microfluidic Mixing Platform

Published: February 25, 2021 doi: 10.3791/62226
Automatic Translation
*1, *1, 1, 1, 1
1Biologics Drug Product Development & Manufacturing, Sanofi, Framingham, Massachusetts
* These authors contributed equally

Summary

Lipid nanoparticles are developed using a microfluidic mixing platform approach for mRNA and DNA encapsulation.

Abstract

Lipid-based drug carriers have been used for clinically and commercially available delivery systems due to their small size, biocompatibility, and high encapsulation efficiency. Use of lipid nanoparticles (LNPs) to encapsulate nucleic acids is advantageous to protect the RNA or DNA from degradation, while also promoting cellular uptake. LNPs often contain multiple lipid components including an ionizable lipid, helper lipid, cholesterol, and polyethylene glycol (PEG) conjugated lipid. LNPs can readily encapsulate nucleic acids due to the ionizable lipid presence, which at low pH is cationic and allows for complexation with negatively charged RNA or DNA. Here LNPs are formed by encapsulating messenger RNA (mRNA) or plasmid DNA (pDNA) using rapid mixing of the lipid components in an organic phase and the nucleic acid component in an aqueous phase. This mixing is performed using a precise microfluidic mixing platform, allowing for nanoparticle self-assembly while maintaining laminar flow. The hydrodynamic size and polydispersity are measured using dynamic light scattering (DLS). The effective surface charge on the LNP is determined by measuring the zeta potential. The encapsulation efficiency is characterized using a fluorescent dye to quantify entrapped nucleic acid. Representative results demonstrate the reproducibility of this method and the influence that different formulation and process parameters have on the developed LNPs.

Introduction

Drug carriers are used to protect and deliver a therapeutic with typical favorable properties including low cytotoxicity, increased bioavailability, and improved stability1,2,3. Polymeric nanoparticles, micelles, and lipid-based particles have previously been explored for nucleic acid encapsulation and delivery4,5,6,7. Lipids have been used in different types of nanocarrier systems, including liposomes, and lipid nanoparticles, as they are biocompatible with high stability8. LNPs can readily encapsulate nucleic acids for gene delivery9,10. They protect the nucleic acid from degradation by serum proteases during systemic circulation11 and can improve delivery to specific sites, as the surface topography and physical properties of LNPs influence their biodistribution12. LNPs also improve tissue penetration and cellular uptake9. Previous studies have demonstrated the success of siRNA encapsulation within an LNP13, including the first commercially available LNP containing siRNA therapeutic for the treatment polyneuropathy of hereditary transthyretin-mediated amyloidosis14 treatment that was approved by United States Food and Drug Administration (FDA) and European Medicines Agency in 2018. More recently, LNPs are being studied for the delivery of larger nucleic acid moieties, namely mRNA and DNA9. As of 2018, there were ~ 22 lipid-based nucleic acid delivery systems undergoing clinical trials14. Additionally, mRNA containing LNPs are currently leading candidates and have been employed for a COVID-19 vaccine15,16. The potential success for these non-viral gene therapies requires forming small (~100 nm), stable, and uniform particles with high encapsulation of the nucleic acid.

Use of an ionizable lipid as a main component in the LNP formulation has shown advantages for complexation, encapsulation, and delivery effciciency14. Ionizable lipids typically have an acid dissociation constant (pKa) < 7; for example, dilinoleylmethyl-4-dimethylaminobutyrate (D-Lin-MC3-DMA), the ionizable lipid used in the FDA approved LNP formulation, has a pKa of 6.4417. At low pH, the amine groups on the ionizable lipid become protonated and positively charged, allowing for the assembly with negatively charged phosphate groups on mRNA and DNA. The ratio of amine, "N", groups to phosphate, "P", groups is used to optimize the assembly. The N/P ratio is dependent on the lipids and nucleic acids used, which varies depending on the formulation18. After formation, the pH can be adjusted to a neutral or physiological pH to allow for therapeutic administration. At these pH values, the ionizable lipid is also deprotonated which imparts neutral surface charge to the LNP.

The ionizable lipid also aids in endosomal escape19,20. LNPs undergo endocytosis during cellular uptake and must be released from the endosome in order to deliver the mRNA cargo into the cell cytoplasm or DNA cargo to the nucleus21. Inside the endosome is typically a more acidic environment than the extracellular medium, which renders the ionizable lipid positively charged22,23. The positively charged ionizable lipid can interact with negative charges on the endosomal lipid membrane, which can cause destabilization of the endosome allowing for the release of the LNP and nucleic acid. Different ionizable lipids are currently being studied for improving efficacy of both LNP distribution, as well as endosomal escape14.

Other typical components of an LNP include helper lipids, such as a phosphatidylcholine (PC) or phosphoethanolamine (PE) lipid. 1,2-Dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE), 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC), and 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) are commonly used helper lipids24,25. DOPE has been shown to form an inverted hexagonal II (HII) phase and enhance transfection by membrane fusion26, while DSPC has been thought to stabilize LNPs with its cylindrical geometry27. Cholesterol is also incorporated in the formulation in order to increase membrane rigidity, subsequently aiding in the stability of the LNP. Finally, lipid-conjugated polyethylene glycol (PEG) is included in the formulation to provide the necessary steric barrier to aid in particle self-assembly27. PEG also improves the storage stability of LNPs by preventing aggregation. Furthermore, PEG is often used as a stealth component and can increase the circulation time for the LNPs. However, this attribute can also pose challenges for recruitment of LNPs to hepatocytes through an endogenous targeting mechanism driven by apolipoprotein E (ApoE)28. Thus, studies have investigated the acyl chain length for diffusion of PEG from the LNP, finding that short lengths (C8-14) dissociate from the LNP and are more amenable to ApoE recruitment compared to longer acyl lengths28. Further, the degree of saturation of the lipid tail that PEG is conjugated to has been shown to influence the tissue distribution of LNPs29. Recently, Tween 20, which is a commonly used surfactant in biological drug product formulations and has a long unsaturated lipid tail, was shown to have high transfection in draining lymph nodes compared to PEG-DSPE, which largely transfected the muscle at the injection site29. This parameter can be optimized to achieve the desired LNP biodistribution.

Conventional methods of forming LNPs include the thin-film hydration method and ethanol-injection method27. While these are readily available techniques, they are also labor intensive, can result in low encapsulation efficiency, and are challenging to scale up27. Advancements in mixing techniques have resulted in methods more amenable to scale up, while developing more uniform particles27. These methods include T-junction mixing, staggered herringbone mixing, and microfluidic hydrodynamic focusing27. Each method has a unique structure, but all allow for rapid mixing of an aqueous phase containing the nucleic acid with an organic phase containing the lipid components, resulting in high encapsulation of the nucleic acid27. In this protocol, rapid and controlled mixing through a microfluidic cartridge is utilized, which employs the staggered herringbone mixing design. This protocol outlines the preparation, assembly, and characterization of nucleic acid containing LNPs.

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Protocol

A schematic of the overall process is provided in Figure 1.

1. Preparation of buffers

NOTE: Sterile filtering of the buffers is highly suggested here to remove any particulates which may impact the nucleic acid and LNP quality.

  1. Phosphate Buffered Saline (PBS)
    1. Prepare 1x PBS using 8 mM Na2HPO4, 2 mM KH2PO4, 137 mM NaCl, and 2.7 mM KCl in nuclease free water and adjust the pH to 7.4.
    2. Sterilize by vacuum filtration using a 0.22 µm pore-size filter.
  2. Citrate Buffer
    1. Prepare citrate buffer using 5 mM sodium citrate, 5 mM citric acid, and 150 mM sodium chloride in nuclease free water and adjust to pH 4.5.
    2. Sterilize by vacuum filtration using a 0.22 µm pore-size filter.
      ​NOTE: Citrate buffer only needs to be prepared if mRNA is the nucleic acid that will be encapsulated in the LNP. If DNA will be encapsulated skip 1.2 and proceed to 1.3.
  3. Malic Acid Buffer
    1. Prepare malic acid buffer using 20 mM malic acid and 30 mM sodium chloride in nuclease free water and adjust to pH 3.0.
    2. Sterilize by vacuum filtration using a 0.22 µm pore-size filter.
      ​NOTE: Malic acid buffer only needs to be prepared if DNA is the nucleic acid that will be encapsulated in the LNP. Skip 1.3 if mRNA is to be encapsulated. Citrate buffer is used for mRNA encapsulation, as the lower pH of 3.0 with malic acid buffer may lead to an increased likelihood of mRNA degradation. The protocol can be paused here.

2. Preparation of lipid mix

  1. If stock lipids are in powder form, solubilize in pure 200 proof ethanol.
  2. Calculate the required mix of lipid components based on the desired molar ratio. A molar ratio of 50:10:39:1 (ionizable lipid:helper lipid:cholesterol:PEG) will be used here as an example for a total lipid concentration of 10 mM. Table 1 shows the concentrations and volumes needed for each of these components.
    NOTE: When calculating the volume needed to achieve the lipid mix concentration in ethanol (EtOH) for the microfluidic mixer, the total volume is accounted for to ensure that the addition of EtOH does not influence the lipid concentrations. For example, an ionizable lipid volume of 68.5 µL is calculated by multiplying the 5 mM concentration in ethanol by a total lipid mix volume of 533 µL and then dividing by the stock lipid concentration of 38.9 mM.
  3. Add the appropriate amount of each lipid stock solution to a glass vial to allow components to mix with intermittent vortexing. Add 200 proof ethanol for a total mixture of 533 µL. For the example in Table 1, this is 254 µL of ethanol.
    ​NOTE: For a single run to produce 1 mL of LNPs, 342.5 µL of lipid solution is needed. This is due to a 3:1 mix of aqueous nucleic acid to organic lipid solution with some volume discarded before and after sample collection. A mix of 533 µL is made to compensate as overage.

3. Preparation of nucleic acid solution

NOTE: Preparation and handling of nucleic acid solutions is to be performed in a sterile and RNase-free environment wherever possible. Work in a biosafety cabinet whenever possible with the nucleic acid.

  1. Calculate N/P ratio. The N/P ratio is the total number of ionizable lipid amine groups (N) to the total number of negatively charged nucleic acid phosphate groups (P). N/P ratio is often a parameter that can be optimized during LNP formation. Follow the steps below.
    1. Calculate the number of N units using the below formula:
      Equation 1
      NOTE: The ionizable lipid concentration (Table 1) is 5 mM, which is equivalent to 5 x 10-6 mol/mL. The required lipid injection volume is 0.3425 mL. For example, if the number of N units per molecule is 1, using the above equation, there are 1.03 x 1018 N units in the lipid mix.
    2. Calculate the P units for the desired N/P ratio. Here an N/P = 36 is used for example.
      Equation 2
  2. Calculate the necessary nucleic acid concentration to obtain 2.86 x 1016 P units using the below equation.
    Equation 3
    Where, the number of P units per base pair for mRNA is 1 and DNA is 2. For an mRNA with 1,200 bases, the amount of mRNA required for a N/P = 36 is 3.96 x 10-11 moles.
  3. Calculate the mass concentration of mRNA required for N/P = 36 using the below equation.
    Equation 4
    The average molecular weight of a ribonucleotide monophosphate unit is 322 g/mol30. With 1,200-base mRNA, the molecular weight of the mRNA is 386,400 g/mol. The required injection volume of nucleic acid solution is 1.028 mL. Thus, the concentration of mRNA needed is 1.488x10-5 g/mL, which is 14.88 µg/mL.
  4. Make up 1.5 mL of 14.88 µg/mL of mRNA in citrate buffer.
    ​NOTE: When DNA will be the nucleic acid encapsulated, use malic acid buffer to make up the nucleic acid solution.

4. Priming the microfluidic channels

NOTE: This protocol is adapted from the instrument manufacturer's guidelines.

  1. Input the priming parameters into the instrument software by clicking on the appropriate fields (Table 2).
    NOTE: A flow ratio of 3:1 and flow rate of 4-12 mL/min is recommended27,31 . This has been shown to be optimal in the studies presented here, as well as by the manufacturer. This can be varied if it is of interest towards the application.
  2. Open the instrument lid and place a microfluidic cartridge into the rotating block.
  3. Draw at least 0.5 mL of ethanol into a 1 mL syringe, ensuring there are no bubbles or air gaps at the syringe tip. Load this syringe into the right inlet of the cartridge.
  4. Fill a 3 mL syringe with 1.5 mL of aqueous buffer (citrate for RNA and malic acid for DNA), ensuring there are no air bubbles or gaps. Load this syringe into the left inlet of the cartridge.
  5. Insert two 15 mL conical tubes in the clip holders to serve as waste containers.
  6. Click on Run in the instrument software to begin the mixing, ensuring that the parameters are input correctly.
  7. When the instrument stops priming, indicated by the bottom blue light shutting off, open the lid and properly dispose of the conical tubes and syringes.

5. LNP formation

NOTE: This protocol is adapted from the instrument manufacturer's guidelines.

  1. Update the software with the formulation parameters by clicking on the appropriate fields (Table 2).
  2. Fill a 1 mL syringe with the lipid mix (prepared in step 2). Remove any air gaps or bubbles at the syringe tip and insert the syringe into the right side of the cartridge.
  3. Draw the nucleic acid solution (prepared in step 3) into a 3 mL syringe, ensuring there are no bubbles or air gaps in the syringe tip. Insert the syringe into the left inlet of the cartridge.
    NOTE: Volumes are provided to make a 1 mL solution of LNPs. This instrument can incorporate syringe sizes up to 10 mL, and volumes can be scaled accordingly with no influence on the outcome. The maximum volume of LNPs that can be prepared in one preparation is 12 mL.
  4. Label a 15 mL RNase-free conical tube with the sample name and insert into the left tube clip. Place a 15 mL waste conical in the right tube clip.
  5. Close the instrument lid and click Run, after confirming correct input of parameters.
  6. After the instrument is finished running, properly discard the waste container and cartridge. Retain the conical tube with the LNP sample.
  7. Dilute the LNP 5x with PBS to minimize the ethanol to <5% (v/v).
    ​NOTE: It is important to dilute the LNPs in PBS as soon as possible after microfluidic mixing to prevent degradation. Always perform the dilution in a biosafety cabinet and continuing to work in the biosafety cabinet throughout the buffer exchanges.

6. Buffer exchange

NOTE: Protocol for using ultra-centrifuge filters is provided. While this method results in a more time efficient exchange of buffers, dialysis may be substituted here.

  1. Pre-wash an ultra-centrifuge filter (100 kDa pore size) with 2 mL of PBS by centrifuging at 1000 x g for 5 min. Empty the PBS from the bottom compartment.
    NOTE: PBS is chosen to increase the pH to 7.4 ± 0.2, which is physiologically relevant and will result in the ionizable lipid having a neutral charge.
  2. Add diluted LNPs to the top compartment of the pre-washed ultra-centrifuge filter and centrifuge at 1000 x g for 12 min.
  3. Discard the flow-through from the bottom compartment. Perform two more washes by adding 5 mL of PBS to the ultra-centrifuge filter each time. Centrifuge at the same parameters. There is no maximum volume that needs to be maintained.
    NOTE: If a scaled-up volume of LNPs were prepared, increase the volume of PBS for each wash accordingly. For example, if 2 mL of LNPs were prepared in a single run, then 10 mL PBS per wash is suggested.
  4. Pipette the LNP solution against the walls of the ultra-centrifuge filter a few times to minimize LNP loss. Remove the LNP solution from the ultra-centrifuge filter and store in a nuclease free vial. Add PBS if needed to achieve a final volume of the LNP solution of 1 mL.
  5. Filter through a pre-wet 0.2 µm syringe filter, if needed.
    ​NOTE: The protocol can be paused here.

7. Measure encapsulation efficiency

  1. Prepare a standard curve by making 2-fold serial dilutions of working nucleic acid solution in PBS, starting with a highest concentration of 500 ng/mL, and making at least five dilutions. Use PBS as a blank.
  2. Prepare the LNP sample dilutions. Dilute LNP samples with PBS, to achieve an approximate theoretical concentration that lies around the mid-point of the standard curve (eg. ~ 250 ng/mL of nucleic acid estimated from the initial concentration).
  3. Prepare a solution of the RNA quantification reagent (for mRNA measurements) with TritonX-100 to disrupt the LNPs and measure the total amount of nucleic acid inside and outside of the LNP. This solution contains 0.5% (v/v) RNA reagent, 0.4% (v/v) TritonX-100, and 99.1% (v/v) PBS.
  4. Prepare a solution of the reagent without TritonX-100 to measure the amount of nucleic acid not encapsulated in the LNPs. This solution contains 0.5% (v/v) RNA reagent and 99.5% (v/v) PBS.
    NOTE: If LNPs encapsulate double stranded DNA (dsDNA), such as plasmid DNA, use the dsDNA reagent in 7.3 and 7.4 instead, following the same procedure.
  5. In a 96-well black fluorescence capable plate, load at least four replicates of each of the LNP and nucleic acid standard solutions prepared in 7.1 and 7.2.
  6. To half of the replicates of standards and samples, add an equal volume of the reagent containing TritonX-100. This will quantify the total amount of nucleic acid.
  7. To the remaining wells of standards and samples, add an equal volume of the reagent without TritonX-100. This will quantify the amount of nucleic acid not encapsulated inside the LNP.
  8. Shake the plate for 5 min at room temperature to ensure thorough mixing of standards and samples with the added reagent, taking precautions to avoid light exposure.
  9. Measure the fluorescence using a microplate reader, with an excitation wavelength of 480 nm and an emission wavelength of 520 nm.
  10. Calculate the concentration of nucleic acid outside of the LNP using the standard curve made with the addition of the reagent without TritonX-100. Multiply by the dilution factor used in 7.2.
  11. Calculate the concentration of nucleic acid both inside and outside of the LNP using the standard curve made with the addition of the reagent containing TritonX-100. Multiply by the dilution factor used in 7.2.
  12. Calculate the concentration of nucleic acid inside by subtracting the concentration of nucleic acid outside (calculated from step 7.10) from the total concentration of nucleic acid both inside and outside (calculated from step 7.11)
  13. Quantify the encapsulation efficiency from the ratio of the concentration of nucleic acid inside the LNP (calculated from step 7.12) and the total concentration of nucleic acid (calculated from step 7.11).
    ​NOTE: The protocol can be paused here.

8. Concentration adjustments

  1. If needed, adjust the nucleic acid concentration within the LNP solution using the results from the encapsulation efficiency.
  2. If a less concentrated solution is desired, dilute the solution with PBS to achieve the desired concentration.
  3. If a more concentrated solution is desired, perform additional centrifugation runs using an ultra-centrifuge filter.
    ​NOTE: The protocol can be paused here.

9. Measure LNP hydrodynamic size and polydispersity

  1. Dilute an aliquot of the LNP solution 40x with PBS to obtain a final volume of 1 mL.
    NOTE: This dilution may be changed if required. This dilution value is suggested as it uses a small volume of the LNP stock solution while providing quality results.
  2. Using a semi-micro cuvette, measure the hydrodynamic diameter and polydispersity index. Add the LNP solution into the cuvette and insert into the instrument. Set up an operating procedure in the instrument software to include the measurement type, sample details (material, dispersant, temperature, and cell type), and measurement instructions (number of runs). Click Start when ready to begin the measurement acquisition.

10. Measure LNP zeta potential

  1. Dilute an aliquot of the LNP solution 40x with nuclease free water to obtain a final volume of 1 mL.
    ​NOTE: Nuclease free water is used as the solvent for zeta potential measurements to minimize the influence of high salt buffers on conductivity.
  2. Using a folded capillary zeta cell, measure the zeta potential.
    1. Add the LNP solution into the cuvette up to the fill line. Insert into the instrument ensuring that the electrodes are making contact with the instrument.
    2. Set up an operating procedure in the instrument software to include the measurement type, sample details (material, dispersant, temperature, and cell type), and measurement instructions (number of runs). Click Start when ready to begin the measurement acquisition.

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Representative Results

Multiple batches of LNPs with the same lipid formulation and N/P ratio of 6 were developed on separate days to demonstrate reproducibility of the technique. Batch 1 and 2 resulted in overlapping size distributions with similar polydispersity (Figure 2A) No significant difference was observed in the size or encapsulation efficiency between the two different batches (Figure 2B). The encapsulation efficiency was high for each batch (>98.5%) and the sizes were similar with a 77 nm LNP diameter. The particles were uniform with an average polydispersity index (PDI) of 0.15 for batch 1 and 0.18 for batch 2.

Changes in formulation parameters showed some small, yet statistically significant differences with respect to the N/P ratio, ionizable lipid used, and nucleic acid encapsulated. While differences are discussed, it is important to note that all LNPs formed resulted in encapsulation greater than 80%, with most formulations greater than 95%, and particle sizes less than 110 nm, making all formulations developed here desirable for gene delivery. First, ionizable lipid A was used to develop LNPs at an N/P of 10 and 36. Decreasing the N/P ratio resulted in a 4% decrease in encapsulation efficiency and an increase in the hydrodynamic diameter of the LNPs from 98 nm at N/P = 36 to 109 nm at N/P = 10 (Figure 3A). Comparing LNPs with ionizable lipid A to a different ionizable lipid B and maintaining N/P of 36 resulted in a significant change in encapsulation efficiency, where 100% of pDNA was encapsulated with LNPs formed using ionizable lipid A and 81% of pDNA was encapsulated with LNPs formed using ionizable lipid B (Figure 3B). Ionizable lipid B LNPs also resulted in slightly smaller particles with a hydrodynamic diameter of 95 nm. Finally, LNPs were formed using ionizable lipid A with both mRNA and pDNA. LNPs encapsulating pDNA resulted in larger particles with a 119 nm diameter compared with mRNA LNPs with a 91 nm diameter (Figure 3C). Both pDNA and mRNA LNPs resulted in similar encapsulation efficiency at ~91-94%.

Lastly, changes in the flow rate process parameter did not impact the LNPs developed at the flow rates tested here. At both 4 mL/min and 12 mL/min, LNPs were developed and characterized to have encapsulated 96% of pDNA and have a 110 nm diameter (Figure 4). All LNPs regardless of process parameter or formulation parameter resulted in charge neutral zeta potential measurements.

Figure 1
Figure 1: LNP development and characterization workflow. First, lipid mix and nucleic acid solutions are made (1 and 2). The lipid mix contains the ionizable lipid, helper lipid, cholesterol, and PEG in ethanol, while the nucleic acid solution contains either mRNA or DNA in buffer. Solutions are mixed using a microfluidic cartridge (3), which forms LNPs (4). Next, a buffer exchange is required to remove the ethanol and increase the solution pH to neutral (5). Characterization of LNPs is performed to determine encapsulation efficiency and particle size, polydispersity, and zeta potential using a fluorescence microplate assay and zetasizer, respectively (6 and 7). Please click here to view a larger version of this figure.

Figure 2
Figure 2: Batch to batch reproducibility of LNPs formed on separate days. (A) Size distributions for batch 1 vs. batch 2 (B) Encapsulation efficiency (%) and hydrodynamic diameter (nm) for each batch with mRNA and N/P = 6. Error bars note standard deviation. Statistical analysis using two-way ANOVA with α = 0.05 shows no significance. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Variations of formulation parameters. (A) LNPs formed at N/P = 10 and 36 both using ionizable lipid A with pDNA. (B) LNPs formed with ionizable lipid A and ionizable lipid B both at N/P = 36 with pDNA. (C) LNPs formed with either mRNA or pDNA both using ionizable lipid C at N/P = 6. Error bars note standard deviation. Statistical analysis was performed using two-way ANOVA with α = 0.05; *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Variations of process parameters. LNPs formed at a flow rate of 4 and 12 mL/min using ionizable lipid A with pDNA at N/P = 10. Error bars note standard deviation. Statistical analysis using two-way ANOVA with α = 0.05 shows no significance. Please click here to view a larger version of this figure.

Lipid Molar Ratio Stock lipid concentration (mM) Concentration in ethanol for lipid mix (mM) Volume (µL)
Ionizable lipid 50 38.9 5 68.5
Helper lipid 10 10 1 53.3
Cholesterol 39 20 3.9 103.9
C-14 PEG 1 1 0.1 53.3
EtOH 254
Total 533

Table 1: Example lipid mix to prepare 1 mL of LNPs. The lipid stock concentrations in ethanol provided have been shown to allow for the lipids to solubilize in ethanol, but other stock concentrations may be utilized and will not affect the outcome as long as the lipid is solubilized. Example concentrations of lipids in ethanol for microfluidic mixing are also provided. These concentrations are based on the molar ratio, which can be varied based on the desired LNP preparation.

Priming Formulation
Volume (mL) 2 1.37
Flow Rate Ratio (Aqueous:EtOH) 3:1 3:1
Total Flow Rate (mL/min) 12 4
Left Syringe Size (mL) 3 3
Right Syringe Size (mL) 1 1
Start Waste Volume (mL) 0.35 0.25
End Waste Volume (mL) 0.05 0.05

Table 2: Microfluidic Mixing Benchtop Instrument Software Priming and LNP Formulation Example Parameters

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Discussion

Reproducibility, speed, and low volume screening are significant advantages of using microfluidic mixing to form LNPs compared to other existing methods (e.g., lipid film hydration and ethanol injection). We have demonstrated the reproducibility of this method with no impact on encapsulation efficiency or particle size observed with different LNP batches. This is an essential criterion for any therapeutic, including LNPs, to become clinically available.

The technique described here employs staggered herringbone microfluidic mixing, which results in LNP formation on the time scale of only a few minutes. This mixing uses chaotic advection which is advantageous for mixing control and shortened time27. This mixer enables the aqueous and organic phases to effectively wrap around each other27. Using the staggered herringbone mixing, previous studies have shown that the particles form at the smallest thermodynamically stable size32, which means that the composition tends to influence the size and polydispersity of the LNPs27,32,33. This was observed in the representative results, where the N/P ratio, ionizable lipid used, and nucleic acid encapsulated were the impacting factors on changes in encapsulation efficiency and particle size. Operating parameters, such as flow rate and mixing ratio can also influence the size above a certain threshold, where afterwards the particle size is at its smallest stable size27,33. No change in encapsulation efficiency or particle size was observed when a flow rate of 4 mL/min vs. 12 mL/min was used. Thus, likely both flow rates are above the threshold that would impact the LNP outcome. The example experiment, and results described above, used lipid A and pDNA. It is possible that different ionizable lipids and nucleic acids could have more influence on LNP characteristics with respect to flow rate. Other types of microfluidic mixing include the T-junction, which uses turbulent flow and the microfluidic hydrodynamic focusing method that is based on convective-diffusive mixing27. Compared to these other types of microfluidic mixing techniques for LNP development, the staggered herringbone mixing enables the combination of three important criteria: rapid mixing, minimizing batch to batch variability, and is commercially available27. All three of the microfluidic mixing methods do allow for higher encapsulation efficiency and controlled size compared to conventional lipid film hydration or ethanol injection methods27.

Finally, the ability to produce low volumes of various LNP formulations at the research & development stage is a significant advantage. One challenge of developing LNPs is the number of variables that can be tested and optimized per formulation to achieve the desired outcome and efficacy. Lipids and nucleic acids can be cost prohibitive to screen, troubleshoot, and modify many formulation parameters (e.g., molar ratios, N/P ratios, process parameters, etc.) to find the most suitable LNP for a given application. While low volumes could be a limitation for producing a final formulation at a large scale, the ability to scale up the technique with larger microfluidic mixing instruments is commercially available.

Critical steps of the protocol start with proper storage of lipid stock solutions at the manufacturer's recommendation. LNPs should then be stored at 2-8 °C until further use. For the nucleic acid preparation, the results presented demonstrate that citrate buffer and malic acid buffer are effective at successfully forming LNPs with high nucleic acid encapsulation34, 35. Other buffers may be used instead if desired. If another buffer is chosen, it is important to maintain the pH below the pKa of the ionizable lipid to ensure that the lipid is cationic and can complex with the nucleic acid. When using the microfluidic mixing instrument, it is important to prime the cartridge prior to LNP formation, not to exceed the the cartridge use as recommended by the manufacturer, and to change the cartridge in between different formulation compositions. The most common flow ratio for formation of the aqueous: organic solution is 3:1; however, this can be changed if needed. The flow rate can also be adjusted as desired. Finally, it is important when working with mRNA to ensure an RNase-free environment throughout the entire process. If the desired size or encapsulation efficiency is not achieved, some places to begin troubleshooting include changing the N/P ratio used or the lipid molar percentages. The instrument process described here uses a benchtop model that has a maximum volume limit of 12 mL, although this process is scalable to larger volumes using different microfluidic mixing models.This process can be adapted to changes in lipid mixtures and nucleic acids for use in developing LNPs for various clinical indications. With this flexibility, numerous future applications can be achieved with LNPs to produce different desired formulations. This technique has also been used for developing other types of nanoparticles, including liposomes and polymeric nanoparticles. With some parameter changes, this method can be used for a variety of nanoparticle formulations.

The protocol detailed here describes a reproducible method for achieving mRNA or DNA encapsulated LNPs. In addition to process parameters, additional considerations can influence the LNP outcome. Previous work has also used similar methods to produce LNPs with various nucleic acids, ionizable lipids, N/P ratios, PEG linker length, etc. These parameters can influence the encapsulation efficiency, size, and charge of the particles. The instrument manufacturer has also noted similar changes depending on these parameters that can be optimized18,36. These parameters can further influence the biodistribution and efficacy of the nucleic acid. For example, studies have investigated hydrocarbon chain lengths (C14, C16, and C18) conjugated to PEG and found that the shorter acyl chain of C14 resulted in higher levels of liver uptake compared to the longer acyl chain, which remained in circulation for a longer period of time28. This protocol allows for the formation, optimization, and testing of LNPs with varied compositions, which makes this a versatile process.

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Disclosures

All authors are employees of Sanofi. The authors declare that they have no conflict of interest or competing financial interests.

Acknowledgments

Thank you to Atul Saluja, Yatin Gokarn, Maria-Teresa Peracchia, Walter Schwenger, and Philip Zakas for their guidance and contributions towards LNP development.

Materials

Name Company Catalog Number Comments
1,2-dimyristoyl-rac-glycero-3-methoxypolyethylene glycol-2000 (C-14 PEG) Avanti Polar Lipids 880151P
10 µl Graduated Filter Tips  (RNase-,DNase-, DNA-free) USA Scientific 1121-3810
1000 µl Graduated Filter Tips (RNase-,DNase-, DNA-free) USA Scientific 1111-2831
20 µl Beveled Filter Tips (RNase-,DNase-, DNA-free) USA Scientific 1120-1810
200 µl Graudated Filter Tips (RNase-,DNase-, DNA-free) USA Scientific 1120-8810
3β-Hydroxy-5-cholestene, 5-Cholesten-3β-ol (Cholesterol) Sigma-Aldrich C8667
BD Slip Tip Sterile Syringes (1 ml syringe) Thermo Fisher Scientific 14-823-434
BD Slip Tip Sterile Syringes (3 ml syringe) Thermo Fisher Scientific 14-823-436
BD Vacutainer General Use Syringe Needles (BD Blunt Fill Needle 18G) Thermo Fisher Scientific 23-021-020
Benchtop Centrifuge Beckman coulter
Black 96 well plates Thermo Fisher Scientific 14-245-177
BrandTech BRAND BIO-CERT RNase-, DNase-, DNA-free microcentrifuge tubes (1.5mL) Thermo Fisher Scientific 14-380-813
Citric Acid Fisher Scientific 02-002-611
Corning 500ml Vacuum Filter/Storage Bottle System, 0.22 um pore Corning 430769
Disposable folded capillary cells Malvern DTS1070
Ethyl Alcohol, Pure 200 proof Sigma-Aldrich 459844
Fisher Brand Semi-Micro Cuvette Thermo Fisher Scientific 14955127
Invitrogen Conical Tubes (15 mL) (DNase-RNase-free) Thermo Fisher Scientific AM12500
MilliporeSigma Amicon Ultra Centrifugal Filter Units Thermo Fisher Scientific UFC901024
NanoAssemblr Benchtop Precision Nanyosystems
Nuclease-free water Thermo Fisher Scientific AM9930
Phosphate Buffered Saline (PBS) Thermo Fisher Scientific AM9624
Quant-iT PicoGreen dsDNA Assay Kit Thermo Fisher Scientific  P7589
Quant-iT RiboGreen RNA Assay Kit Thermo Fisher Scientific R11490
Sodium Chloride Fisher Scientific 02-004-036
Sodium Citrate, Dihydrate, granular Fisher Scientific 02-004-056
SpectraMax i3x Molecular Devices
Zetasizer Nano Malvern

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References

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Tags

Lipid Nanoparticles Gene Delivery Microfluidic Mixing Platform Formulation Parameters Bio Distribution Herringbone Design Laminar Flow Scaling Up Encapsulated Particles Gene Therapy Non-viral Approach Repeat Dosing Liquid Stock Solution Glass Vial Vortexing Ethanol MRNA Stock Citrate Buffer Microfluidic Channels Priming Parameter Instrument Software Rotating Block
Formulating and Characterizing Lipid Nanoparticles for Gene Delivery using a Microfluidic Mixing Platform
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Bailey-Hytholt, C. M., Ghosh, P.,… More

Bailey-Hytholt, C. M., Ghosh, P., Dugas, J., Zarraga, I. E., Bandekar, A. Formulating and Characterizing Lipid Nanoparticles for Gene Delivery using a Microfluidic Mixing Platform. J. Vis. Exp. (168), e62226, doi:10.3791/62226 (2021).

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