By George Kamphaus, Ph. D.

Hydrogen fuel cells are already used as “clean” energy sources. Hydrogen can be produced using a number of different processes: Thermochemical processes that use organic materials, Electrolytic and Photolytic processes that split water (H₂O) into hydrogen (H₂) and oxygen (O₂), and a more recently introduced method in which microorganisms such as bacteria and algae can produce hydrogen through biological processes. This blog discusses the use of genetically modified purple non-sulfur photosynthetic bacterium Rubrivivax gelatinosus CBS as a new platform for biological H₂ production developed by C. Eckert, et al. at the National Renewable Energy Center and the University of Colorado.

• Electroporation ready cells of CBS were made by growing cells photosynthetically in 14 ml RCVBN media with sodium malate to mid-logarithmic phase of growth (optical density [OD₆₀₀] between 1 and 2). Cells were then harvested by centrifugation (6600 x g for 5 minutes), washed twice with cold ddH2O and resuspended in 1 ml of 10% glycerol. This suspension was separated into 80 μl aliquots and stored at -80°C.
• Cell aliquots (80 μl) were mixed with 1 g of the purified integration vectors in chilled 1 mm gap cuvettes (BTX Harvard Apparatus).
• The BTX ECM 630 was set to 2.0 kV, 200 Ω, 25 μF, and one pulse. Cuvettes were pulsed with a time constant of ~5 ms.
• Cells were diluted into 900 μl RCVBN media with sodium malate in a 2 ml microcentrifuge tube and were incubated in the dark on a nutator at 30°C. 100 μl of this outgrowth was plated onto agar plates of RCVBN media with sodium malate plus 50 μg/ml kanamycin and/or 100 μg/ml carbenicillin and incubated in the dark at 30°C.

• R. gelatinosus CBS, unlike related bacteria, can metabolize CO to produce H₂ in a “water-gas shift” reaction (CO + H₂O → H₂ + CO₂)
•  Prior studies by these authors using transposon mutagenesis and whole genome sequencing have identified numerous putative genes involved in the CO oxidation reaction and CO-linked H₂ production pathway as well as CO-sensing transcription factors.
• This 2019 report discovered two clusters of hyp maturation factors (hyp 1 and hyp 2) through homology comparison using the whole genome sequence. The hyp maturation factor proteins A – F are critical for the assembly of the complex multiunit hydrogenases in many microorganisms.
• This paper characterized the induction of the hyp1 and hyp2 genes in the presence of CO and or H₂ and determined hyp1 factors were clearly induced in the presence of CO but not in the presence of H₂.
• Addition of CO to the bacteria culture also induced the cooH gene, which encodes the catalytic subunit of the Ech (energy-converting hydrogenase). This indicates that the hyp1 genes are maturation factors involved in the Ech assembly. H₂, independent of CO, was able to induce all hyp2 genes tested as well as membrane bound hydrogenase (MBH) gene hupB. 
• While it is also true that CO addition also increased hyp2 and hupB transcripts, this is likely due to the production of H₂ from CO metabolism. Taken together, this supports the conclusion that hyp2 factors are critical to the assembly (or maturation) of MBH hydrogenases.

• All mutant strains (hyp1 Δ , hyp2 Δ  and hyp1/2 Δ) grew similarly to wild type following CO addition under photosynthetic, mixotrophic conditions in medium containing malate and yeast extract as additional carbon sources over the course of the 24 hour experiment, as determined by OD₆₀₀ readings.
• Hyp2 Δ mutants were able to metabolize added CO at similar rates as wild-type cultures. Hyp1 Δ  mutants showed slower metabolism of CO and hyp1/2 Δ double knockouts showed almost no CO metabolism indicating that the while hyp1 factors are likely maturation factors of the Ech hydrogenase complex. It is possible that the hyp2 factors can partially compensate for the hyp 1 factors that were deleted.
• While both wild-type and hyp1 Δ mutants produce H₂ in response to added CO (peaks at 8 hours and 12 hours respectively) both of these cultures metabolized the H₂, likely due to oxidation by the MBH hydrogenase to produce energy for the cell.  The hyp1/2 Δ mutants showed no H₂ production, while the hyp2Δ mutants produced levels of H₂ nearly 3.5 times the peak of the wild-type cells and which was sustained for the duration of the experiment.
• To further test the theory that hyp2 genes are involved in H₂ metabolism, H₂ gas was injected into the “headspace” of cultures and the concentration of the gas was monitored for 24 hours. Both wild-type and hyp1Δ mutants depleted the H₂ level to zero by 16 hours, while both the hyp2Δ  and hyp1/2 Δ mutants showed limited hydrogen depletion.
• Western blot analysis of Ech hydrogenase apoprotein indicated no major differences in expression among the mutants compared to wild-type bacteria. This indicates that the observed changes in H₂ production of hyp1Δ mutants at the 8 hour time point is likely due to delayed assembly of the CooH active site rescued by Hyp2 in the absence of Hyp1 proteins.

Reference:

1. Eckert, C. A., et al.  (2019) Inactivation of the uptake hydrogenase in the purple non-sulfur photosynthetic bacterium Rubrivivax gelatinosus CBS enables a biological water–gas shift platform for H2 production. Journal of Industrial Microbiology and Biotechnology, 46(7), 993-1002.

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By Michelle M. Ng-Almada, Ph. D.

In this blog we will first overview the model system Chlamydomonas reinhardtii and methods for transforming this single celled, flagellated alga.  We will provide protocol recommendations for high-efficiency square wave electroporation with generators such as the ECM 830 which can achieve a transformation efficiency of 2 to 6 x 10³ transformants per µg of exogenous DNA.  Next, we will cover protocol recommendations for exponential decay wave generators, such as the ECM 630, which can achieve transformation efficiencies of 2 to 3 x 10³ transformants per µg of exogenous DNA.  Finally, we will compare the pros and cons and typical efficiencies of these methods to traditional glass bead transformation and electroporation via decaying square wave generators.

Chlamydomonas reinhardtii is an excellent model system for studying basic biological structures and processes

• Chlamydomonas has two flagella which are similar in structure and function to cilia in other organisms.  These flagella can be isolated for biochemical and molecular analysis, and have been used for many years as a model system to study cilia biogenesis and intraflagellar transport (IFT).
• Chlamydomonas may be used to study photosynthesis. It can grow on simple salt medium, utilizing photosynthesis to derive energy, or can grow in comlet darkness if acetate is provided as an alternate carbon source.
• C. reinhardtii exists in a haploid vegetative state, or under adverse environmental conditions the two mating types may fuse and create a diploid zygospore.  When the conditions improve, the diploid zygote undergoes meiosis and releases four haploid cells that resume the vegetative life cycle.
• The full sequences of the nuclear, chloroplast, and mitochondrial genomes of C. reinhartdii have been sequenced and mapped.
• These characteristics allow for both forward and reverse genetic screening techniques.
• Additionally, the Chlamydomonas Resource Center offers a repository for wild type and mutant cultures and reagents.
• Electroporation is an efficient, convenient method of introducing exogenous DNA without having to remove the Chlamydomonas cell wall.

High-Efficiency Square Wave Electroporation Protocol

• Grow cells in liquid TAP medium to a cell density of 1 to 2 x 10⁷ cells/ml.
• Inoculate cell stock into fresh liquid TAP medium to a concentration of 1 x 10⁶ cells/ml and grow under continuous light for 18 to 20 hours until a cell density of 4 x 10⁶ cells/ml is reached.
• Centrifuge cells at 1250 g for 5 minutes at room temperature, wash, and resuspend with pre-chilled TAP medium containing 60 mM sorbitol, and put the mixture on ice for 10 minutes.
• Place 250 µl of cell suspension (or 5 x 10⁷ cells) into a pre-chilled 4 mm gap electroporation cuvette, and add 100 ng of desired exogenous DNA.
• Electroporate with a generator with square wave form, such as ECM 830 or Gemini X2, using the following electroporation parameters: 500 V, 4 ms pulse duration, 6 to 7 pulses, 100 ms pulse interval time.
• Immediately place the cuvette on ice for 10 minutes after electroporation, then transfer cell suspension into a 50 ml conical centrifuge tube containing 10 ml TAP medium to recover overnight with dim light and slow shaking.
• Plate cells as desired with appropriate selection medium.
• Expected transformation efficiency is 2 to 6 x 10³ transformants per µg of exogenous DNA.
• Please see Wang L. et al. 2019¹ for protocol details.

Exponential Decay Wave Electroporation Protocol

• Prepare cells pre- and post- electroporation as described above for Square Wave Electroporation Protocol
• Electroporate with a generator that has exponential decay waveform, such as ECM 630 or Gemini X2, using the following electroporation parameters: 800 V, 1575 Ω resistance, 50 µF capacitance.
• Desired pulse length is 10 to 14 ms; above or below this desired pulse length range may reduce transformation efficiency.
• When tested side by side with above high-efficiency square wave electroporation, this exponential decay wave method yielded approximately 50% of the efficiency of square wave.
• Please see Wang L., et al. 2013² for protocol details.

Comparison with other Chlamydomonas transformation methods

• Glass bead transformation^3,4 utilizes agitation with glass beads to introduce exogenous DNA.  This method does not require special equipment, however it requires the use of cell-wall deficient strains for removal of cell wall prior to the glass bead agitation.  When compared side by side with the above high-efficiency square wave electroporation method, glass bead method was only about 20% of the efficiency of the square wave method.
• Electroporation with another type of electroporator (decaying square wave pulse generator NEPA21), like the square wave and exponential decay wave electroporation methods described above, has the benefit not requiring cell wall removal. Transformation with this type of electroporator is reported to yield 0.4 to 3 x 10³ transformants per µg of DNA⁵. However this method requires a relatively large amount of exogenous DNA (400 µg per reaction) which is not ideal for genetic screening applications.
• In comparison, high-efficiency square wave electroporation and optimized exponential decay wave electroporation with generators such as ECM 830 and ECM 630 yield transformation efficiencies of up to 6 x 10³ transformants per µg of exogenous DNA, does not require removal of cell walls, and requires only 100 µg of exogenous DNA per transformation reaction.

References:
1. Wang, L., et al. (2019) Rapid and high efficiency transformation of Chlamydomonas reinhardtii by square-wave electroporation. Biosci Rep 39 (1): BSR20181210.

2. Wang, L., et al. (2013) Flagellar regeneration requires cytoplasmic microtubule depolymerization and kinesin-13. J. Cell Sci. 126, 1531–1540.

3. Kindle, K.L. (1990) High-frequency nuclear transformation of Chlamydomonas reinhardtii. Proc. Natl. Acad. Sci. USA 87, 1228–1232.

4. Nelson J.A. and Lefebvre P.A. (1995) Transformation of Chlamydomonas reinhardtii. Methods Cell Biol. 47, 513–51.

5. Yamano T., Iguchi H. and Fukuzawa H. (2013) Rapid transformation of Chlamydomonas reinhardtii without cell-wall removal. J. Biosci. Bioeng. 115, 691–694. 

Buy an ECM 830 Electroporation System, get a free BTXpress High Performance Electroporation Buffer—Click here for promotion details!

By George Kamphaus, Ph. D.

This blog describes a new method for creating vaccines against tumors. Shadi-Yunger, et al.¹ have created a novel MHC-II platform by generating a chimeric invariant chain, in which the semi-peptide CLIP is replaced with essentially any tumor specific peptide. This hybrid MHC-II chain facilitates the activation of CD4+ T-cells when expressed in dendritic cells (DCs).  In this particular study, the researchers attempted to create an anti-melanoma vaccine by inserting different melanoma-associated antigens (MAAs) mRNA sequences into the MHC-II hybrid construct. The hybrid MHC-II chain is then combined with another novel MHC-I light chain chimera created by the same group. The MHC-I construct converts the light chain into an integral membrane protein by linking an antigenic peptide (in this case an MAA peptide sequence) at the extracellular N-terminus and coupling to either the membrane anchoring K^b sequence or the intracellular TLR4 signaling domain at the C-terminus. In multiple experiments, different MAA sequences were inserted into the hybrid constructs and mRNAs coding both the hybrid MHC-I and MHC-II were transfected into murine bone marrow derived dendritic cells (BMDCs) by electroporation using the BTX ECM 830 generator. These transfected BMDCs were then injected into mice bearing tumors from Ret melanoma cells, to assess the anti-tumor activity and T-cell activation stimulated by these modified DCs.

A brief description of the electroporation protocol (Please see Shadi-Yunger et al.¹ for method details.)

• BDMCs isolated from femurs and tibiae of 4 to 5 weeks old C57BJ/6 female mice were cultured for 10 days. On the day of transfection cells were washed two times with Opti-MEM (GibcoBRL) and resuspended into Opti-MEM medium containing 10 to 20 g of transcribed mRNA.
• Add mixture to electroporation cuvette (for example, Cuvettes Plus, 2 mm gap, BTX).
• Electroporation: 1 pulse (square wave) at 400 V, 0.9 ms pulse duration.
• Collect cells from the cuvette and resuspend in 5 ml of pre-warmed media and place into 50 ml tube for further incubation.

Selecting MAAs for the anti-tumor vaccine

Since prior studies had shown spontaneous anti-melanoma immune responses could be directed against melanocyte differentiation antigens such as gp100, tyrosinase (Tyr) and tyrosinase related proteins 1 and 2 (TRP-1 and TRP-2), these proteins have been studied to find peptides or fragments that could enhance immune anti-tumor responses. Prior to this study, the authors had tested the anti-tumor activity of two such peptides as part of chimeric polypeptide sequences, gp100_25-33 and TRP2_180-188. By further expanding their search using SYFPEITHI prediction software, they were able to find several other peptides for study as MHC-I MAAs. The two chosen for hybridization in the MHC-I constructs this study were TRP1_455-463 (designated E120 in the figures) and Tyr_360-368 (called E124) both of which bind, or are predicted to bind to H-2D^b  

Two different MAA peptides were chosen for the creation of MHC-II chimeric proteins. The first was TRP-1_111-128 (designated E122) which was previously shown to confer immune response, and Tyr_99-117 (aka E130) which was predicted to induce an immune response by the IEDB prediction tool.

Characterization and functional analysis of transfected BMDCs

• Cells were assessed for expression of MHC-I chimeric constructs by flow cytometry 6 hrs after electroporation using anti-β2m antibody.
• MHC-II constructs could not be assessed using commercial antibodies, so instead surface expression was determined by the ability to induce proliferation in T-cells from mice immunized with the corresponding MAA peptide. Proliferation was assessed by in vitro H³-thymidine uptake assays. Two different hybrid MAA-CLIP MHC-II constructs were able to stimulate CD4+ T-cell proliferation with similar kinetics as the corresponding peptide-loaded BMDCs.
• BMDCs transfected with the hybrid MHC-I constructs alone were able to induce targeted cell killing by T-cells in both in vivo and in vitro CTL experiments.

In vivo anti-tumor vaccine studies

Once the authors concluded that the chimeric MHC-I and MHC-II proteins were expressed and functional when the mRNA constructs were transfected into BMDCs, they performed several in vivo studies in Ret melanoma injected mouse tumor models. The results, summarized below, indicate that a melanoma vaccine will likely require a combination approach; multiple antigens and induction of both MHC-I and MHC-II responses. Furthermore, it appears that CD4+ T-cells are important mediators of anti-tumor response. 

Reference:
1. Sharbi-Yunger, A., et al. (2019). A universal anti-cancer vaccine: Chimeric invariant chain potentiates the inhibition of melanoma progression and the improvement of survival. International Journal of Cancer, 144, 909-921.

Buy an ECM 830 Electroporation System, get a free BTXpress High Performance Electroporation Buffer—Click here for promotion details!

By Michelle M. Ng, Ph. D.

Regeneration of nerve cell axons is required for recovery of function after injury to the nervous system.  A more thorough study of the molecular and cellular mechanisms of this process is needed to better understand this process of axon regeneration.  Previously, quantification of axon regrowth was commonly accomplished with indirect methods of measuring length, such as immunostaining of longitudinal nerve sections. In this post we will review a new method to directly trace regenerating axons in vivo, recently published by Yan Gao, et al. in Neural Regeneration Research.¹ The researchers performed in vivo electroporation of mouse adult sensory neurons in the ipsilateral dorsal root ganglion to transfect plasmid DNA encoding enhanced green fluorescent protein (eGFP).  Next, the sciatic nerve was squeezed with tweezers to create a model of sciatic nerve compression, and finally a direct time course of regenerating axon lengths was captured by confocal microscopy.

Overview of the electroporation protocol (please see Saijilafu et al., 2011² for method details)

• 8- to 10-week-old male CF-1 mice either underwent a sham operation or a sciatic nerve crush combined with electroporation.
• Lumbar 4 and 5 dorsal root ganglion neurons (DRGs) on one side of the mouse were exposed after anesthetization.
• 1 µl of transfectant solution (either 2 to 3 µg of pCMV-EGFP-N1 plasmid DNA or 50 pmol Dy547-tagged microRNA hairpin inhibitor) was injected into DRGs using a capillary pipette.
• Electroporation was then performed using a tweezer-shaped electrode and the BTX ECM 830 Electroporator.
• Electroporator settings were square waveform, 35 volts, 5 pulses, 15 ms duration, 950 ms interval.
• At 2 or 3 days after in vivo electroporation, the sciatic nerve on the side with electroporated DRGs was exposed and crushed at the sciatic notch.
• For the timecourse study, at desired timepoints (12 hours, 18 hours, and 1, 2, 3, 4, 5, and 6 days) the mice were sacrificed, the tissue was perfused and cleared, and the axons were imaged by fluorescence confocal microscopy.

Quantifying axon regrowth after nerve crush

Representative images of sciatic nerves are shown in the left panels below, and quantification is shown in the panel at the below right.  The results of this study showed that sensory axons regenerated across the crush site (red line) very slowly on the first day, then more quickly and steadily by day 2 and beyond.

High-resolution 3D imaging of DRGs and sensory axons

A benzyl alcohol/benzyl benzoate clearing approach was employed by the group to image whole mount DRGs and sensory axons with high resolution. Maximum projection images are shown in the left panels below, and reconstructed 3D images are shown in the right panels below. Transfection efficiency of small RNA oligos (red) was higher than that of larger eGFP plasmids.

Tissue clearing and 3D imaging of whole mount DRG electroporated with EGFP plasmid (A and B) and fluorescent dye-tagged microRNA inhibitor (C and D). Adapted from Figure 3 of Gao et al. 2020

In summary, electroporation mediated labeling of dorsal root ganglion enables the direct study of axon regeneration in vivo.   The researchers were successful in using electroporation to transfect fluorescent expression plasmids or dye labeled miRNAs, and then conducted direct timecourse experiments to directly measure axon regrowth lengths. This electroporation method previously was found to not cause any observable signs of tissue damage or cell abnormalities, and additionally in this study, immunostaining for caspase-3 (a marker of apoptosis) found no difference between control and electroporated groups. 

References:
1. Gao, Y., et al. (2020). Time course analysis of sensory axon regeneration in vivo by directly tracing regenerating axons. Neural Regeneration Research, 15(6), 1160-1165.
2. Saijilafu, H., et al. (2011). Genetic dissection of axon regeneration via in vivo electroporation of adult mouse sensory neurons. Nature Communications, 2, 543.

Buy an ECM 830 Electroporation System, get a free BTXpress High Performance Electroporation Buffer—Click here for promotion details!

This blog describes a novel system for the study of T-cell activation by antigen-presenting dendritic cells (DC) and the use of a BTX ECM 830 electroporator to transfect quiescent (naïve or memory) T-cells. While there are many T-cell therapies that use genetically modified or transfected T-cells, in nearly all cases, the T-cells are activated before transfection. Such methods preclude the study of the physiologic mechanisms of activation by DC cells. To increase the understanding of the intercellular dynamics of quiescent CD8+ T-cell interaction with DC and CD-4+ helper cells, Abu Shah, et. al. have created a 3-D system that reconstitutes a tissue-like environment that allows for the integration of cytokines, DC and quiescent T-cells. Utilizing electroporation to transfect mRNA constructs of modified antigen-specific TCRs, the researchers were able to express functional TCR on the surface of the quiescent T-cells and show activation by interaction with DCs loaded with the corresponding peptide antigen.

• Harvest and wash T-cells three times with Opti-MEM (Thermo Fisher Scientific). Resuspend cells at 25 x 10⁶ cells/ml.
• Aliquot 100 to 200 µl (2.5 to 5 x 10⁶ cells) into separate tubes and mix with the desired mRNA products.
• For 10⁶ CD8 T-cells, use 2 µg of each TCRα, TCRβ and CD3ζ RNA. For 10⁶ CD4 T-cells, use 4 µg of TCRα, TCRβ RNA.
• Add mixture to electroporation cuvette (Cuvette Plus, 2 mm gap, BTX).
• Electroporation Parameters for the BTX ECM 830: 1 pulse (square wave) at 300 V, 2 ms pulse duration.
• Collect cells from the cuvette and culture in 1 ml of pre-warmed media.

• Labeling with different fluorescent markers allows different cell types to be monitored simultaneously by time-lapse video. In this study the motility of CD4+ and CD8+ T-cells and DC were all monitored together in a single system under identical conditions.
• Different chemokines can be added to the matrix.
• T-cells can be added before or after solidification of the collagen matrix to study varying kinetics of the DC / T-cell presentation.
• The effect of TCR-antigen affinity can be studied by expression of various TCR in quiescent T-cells.
• This can be used as a model system for immune-modulation therapies, such as PD-1 checkpoint inhibitors.
• Because cells can be easily extracted after observation, cell surface markers and other characteristics can be studied by FACS or other flow cytometry methods.

• Using the BTX ECM 830 square wave electroporator, the efficiency of TCR expression varied among donors with an average value of 81 ± 7% (mean ± SD, n = 13), with more than 90% cell viability and 80–90% cell recovery, both of which were severely reduced using alternative electroporation approaches (Amaxa and Neon).
• The induction of expression of an exogenous TCR in CD4 T-cells is considerably harder than in CD8 T-cells (Dai et al., 2009). Using natural sequences or the introduction of cysteine modifications used for 1G4 and 868 failed to induce expression of TCRs in naïve CD4 T-cells.
• Cell motility of both CD4+ and CD8+ T-cells was faster than DC, either with or without homeostatic chemokines.
• While both high affinity and low affinity peptide-TCR interactions were able to activate CD8+ T-cells, as evidenced by CD69 and CD25 expression, only T-cells with high-affinity TCR-antigen interactions showed arrested motility.
• Two different anti-PD-1 antibodies enhanced the clustering of CD-8 T-cells around their targets, while not changing the motility dynamics.
• Using the BTX ECM 830 square wave electroporator, the cells maintained the motile behavior and interaction dynamics of naïve CD8 T-cells expressing 1G4, at similar levels to those of untouched cells, on 2D stimulatory ‘spots.'

In honor of National Beer Day, April 7th, the focus of this blog post is how to use electroporation for genetic engineering of brewing yeast. First, we will provide the ideal electroporation parameters for Saccharomyces cerevisiae using either ECM 630 or Gemini X2 electroporators.  Then, we will highlight an example from the primary literature where this electroporation method was used with brewing yeast Saccharomyces cerevisiae ssp. carlsbergensis to modify the amount of ethanol in beer.

1. Grow Saccharomyces cerevisiae up to log phase in YPD media
2. Cetrifuge and wash pelleted cells several times to remove cell culture media and suspend cells in electroporation buffer at desired density of 1.6 x 10⁹ cells/ml
3. Electroporate cells in 2 mm gap cuvettes or 2 mm gap High Throughput electroporation plates with the following conditions:
• Temperature: 4°C
• Voltage: 2500 V
• Capacitance: 25 uF
• Resistance: 100 to 200 Ω
• Number of Pulses: 1
• Desired Electrical Field Strength: 12.5 kV/cm
• Desired Time Constant: 3 to 4.5 ms
4. Allow cells to incubate at 30C for 1 hour in recovery media made up of 1:1 1M sorbitol:YPD media.
5. Proceed to desired selection method.

 Electrofusion Basics - 

Determining Cell Fusion Efficiency

By Michelle M. Ng, Ph. D.

There are different ways of analyzing efficiency following an electrofusion experiment. These methods vary in terms of how soon after electrofusion you may analyze results, what types of equipment/reagents are required, and what type of information is determined.

In this post we will provide an overview of a couple of analysis methods that provide some initial results just minutes after electrofusion. 

Typically after an electrofusion experiment, the gold standard method of assessing fusion efficiency is to dilute and plate the cells, allow for colonies to grow out, and then screen for colonies that exhibit the desired traits of the two parental cell types.

 

For example, in Hybridoma/monoclonal antibody production, fusion products of antibody-producing B-lymphocytes and myeloma cells undergo dilution, plating and selection by growth in HAT media for one to two weeks. HAT media supports the growth of the desired fused hybridoma cells.  Next, colony counting and screening are done one to two weeks after electrofusion. The number of colonies, and more specifically the number of colonies secreting antibodies specific to the antigen of interest indicate the efficiency of the hybridoma fusion reaction.

 

There are, however, a couple of simple cell staining techniques that allow you view an aliquot of fused cells as soon as 30 minutes after electrofusion.

 

1. Nuclear Staining allows a researcher to see fused cells as evidenced by more than one nucleus per cytoplasm. For this, stain a sample of cells post-electrofusion with your desired nuclear stain. Then, visualize the cells on a microscope and check the percentage of cells that are multinucleate. The number of multinucleate cells is proportional to the efficiency of the fusion reaction; however, this method does not specifically provide information regarding how many of these multinucleate cells are the desired fusion product containing both parental cell types. Wright-Giemsa nuclear staining is a simpler method that can be done using a light microscope.

Fusion product of an immune cell and tumor cell stained with Wright-Giemsa 45 minutes after electrofusion.

 

You can find a protocol for Wright-Giemsa nuclear staining here.

2. Cell tracing dyes enable a researcher to specifically monitor the two parental populations of cells and their fusion products post-fusion. For this, the parental cell types may be labeled with two separate vital stains of different fluorescence spectra (such as green and red) prior to fusion. Here is an example publication where the researchers used cell tracker dyes to monitor electrofusion efficiency when fusing myeloma cells with CHO cells. This method allows you to determine how many fused cells originate from Parent A (red) + Parent B (green). The desired hybrid cells (containing both green and red) counted by this method will give you roughly one-half of the fusion efficiency numbers as the nonspecific nuclear staining method. The fluorescent staining method requires some additional cell handling pre-fusion and requires access to a fluorescence microscopy set up with at least two channels for fluorescence. 

These cell staining methods are a great way to get an early measure of the success of an electrofusion experiment. They allow the researcher to compare to other experiments and proceed with confidence to clone selection and screening.

 

Here is a link to an example electrofusion protocol from the BTX protocol database for hybridoma electrofusion applications.

Click here to download our Hybridoma Application Note.

In this post we will highlight a high efficiency method for primary human T cell transfection utilizing electroporation of mRNA.  This method creates genetically engineered T-cells which specifically recognize antigens of interest for CAR T-Cell immunotherapy applications.

Combining Wave Forms to Create an Electrofusion Protocol

By Michelle M. Ng, Ph. D.

The joining of membranes of neighboring cells by the application of a pulsed electrical field is called Electrofusion. Electrofusion is widely used for hybridoma creation for monoclonal antibody production, transgenics/somatic cell nuclear transfer applications, hybrid plant creation and functional studies combining two different cell types with different properties. 

In this post we’ll explore the types of electrical wave forms used for electrofusion and outline the way that these waveforms are combined to make a successful electrofusion protocol.

The process of electrofusion utilizes the properties of two types of electrical wave forms:

 

1) An Alternating Current (AC) Wave Form has electrical current and oscillates in voltage and polarity at a defined frequency.

 AC sine wave form

2) Direct Current (DC) Square Wave pulse has a single polarity and voltage for a defined pulse length time.

DC square wave form

 
A successful electrofusion protocol is made up of the three following stages:
1) Alignment. AC waveform is applied to align cells by a process called dielectrophoresis. The alternating current electric field induces cells to move and act as dipoles with negatively charged and positively charged sides. As cells move the dipoles attract, and the cells line up into pearl chain formations as in the micrograph below.

 

  Pearl chains of aligned cells

 

Once cells are aligned, the electrical field causes adjacent cells to compress together. Electrofusion protocols may include multiple alignment protocol segments of different voltages, frequencies and times, (typically increasing in voltage).

2) Fusion. DC square wave form pulse(s) are applied forming temporary pores in the cell membranes using the same process as in electroporation.  In addition to forming temporary pores in the cells, these pulses create pathways between the cell membranes of adjacent cells.

3) Post-fusion. An AC wave form pulse is applied to hold adjacent cells together while the pathways between fused cells mature.  The waveform holds cells in place with gentle force, similar to the alignment phase, to promote fusion, (typically decreasing in voltage).

 

Here is a link to an example electrofusion protocol from the BTX protocol database for hybridoma electrofusion applications.  It includes a couple of pre-fusion AC steps with increasing voltages, a single DC square wave fusion pulse, and a post-fusion AC step with decreasing voltage.

Click here to visit our Electroporation Education Resource, for more basics.

Transfection of Human Cerebral Organoids

By Michelle M. Ng, Ph. D.

In this post we will review what organoids are and how they are used to create in vitro models to study organ function and human disease states in three dimensions. Next, we will cover the topic of organoid transfection via electroporation in more detail. Finally, as an example, we will highlight and summarize findings reported by Ogawa et al. 2018. Cell Reports 23, 1220—1229¹. These researchers utilized electroporation of CRISPR/Cas9 constructs to achieve gene modification of human cerebral organoids, resulting in a glioblastoma tumor model that could be further studied in vivo in xenografted mice.

Organoids are multicellular cultures of stem cells or tissue fragments grown in vitro, organized into 3D structures. This typically requires growing and differentiating stem cells or progenitor cells in 3D culture media (such as Matrigel). Unlike cultured cell lines, organoids may contain multiple different types of cells and offer a way to better replicate the more complex structures found in real organs in vivo. Organoids have been developed to model many different types of organs, such as kidney, lung, liver, heart, retina, pancreas, intestine and brain.

Organoids allow a researcher to study a simplified miniature version of an organ that recapitulates organ structure and functions more realistically than 2D cultured cell lines adhered to dishes.  For example, it has recently been reported by Trujillo et al. ² that cerebral organoids generated from induced pluripotent stem cells (iPSCs) can be cultured for months or even years in the laboratory.  Over time these cerebral organoids mature into complex networks of different types of neurons and glial cells that can produce brain waves with characteristics similar to that of pre-term human fetuses.Human organoids may also be implanted into laboratory animals such as mice to be studied as orthotopic xenografts in vivo.Sugimoto et al.³ utilized this method to grow human colonic epithelial organoids in orthotopically xenografted mouse colons.

• The 3D structure of organoids makes it difficult to efficiently penetrate cells in the center with chemical, lipid or viral methods.  Electroporation more effectively transfects thicker structures such as organoids, ex vivo cultured pieces of tissue, or even tissues of a live organism in vivo.
• Smaller organoids may be electroporated while suspended in a solution containing the transfectant inside an electrode such as an electroporation cuvette, microslide chamber or petri dish electrode chamber.  For this type of electroporation method, using a high-performance electroporation buffer can boost efficiency.  For example, Fukii et al.⁴ found that electroporation was more efficient in transfecting organoids than reagent-based transfection methods. They also showed that utilizing BTXpress electroporation solution further boosted electroporation efficiency, as compared to electroporation in cell culture medium.
• Larger organoids may be injected with the transfectant, then grasped by paddle-shaped electrodes or tweezer-shaped electrodes for electroporation.  This method was utilized successfully by Ogawa et al.¹ as described in more detail below.

Glioblastoma Model via Electroporation of Human Cerebral Organoids

3. Transfected cells in the organoid were observed to have a transduced, proliferative phenotype characteristic of glioma tumor cells; the HRasG12V-tdTomato expressing cells were observed to invade and take over the organoid over time as shown in the figure below. Organoid tumor cells also were positive when stained for expression of Ki-67 proliferative marker and SOX 2 and GFAP markers of glioblastoma neural stem cells.

Time-lapse imaging of HRasG12V-transduced cells showing invasion (from 4 weeks to 13 weeks post transfection). (a) High-magnification view of the invasive edges of a 13-week transduced organoid. Adapted from Fig 1D of Ogawa et al. 2018.

4. Finally, the researchers injected organoid-derived tumor cells into the hippocampus of immunodeficient mice to observe invasive tumor growth in vivo. They observed tumors forming with pathological features similar to human glioblastoma, such as microscopic invasiveness, nuclear pleomorphism, and angiogenesis. The mean survival times for the mice were 90 to 100 days.  The figure below shows a representative tumor in one of the mice.

Gross appearance and fluorescence signal of xenografted mouse brain. The dotted area shows a slightly pinkish appearance from tdTomato expression. Right panel indicates the fluorescence signal with a red fluorescent protein (RFP) filter. Adapted from Fig 3E of Ogawa et al. 2018.

In summary, electroporation mediated transfection of organoids offers a powerful platform to study simplified, engineered models of organs in vitro.  When used in combination with CRISPR/Cas9 gene editing methods, genetic manipulation of these model organs and their study in vitro is possible. Additionally, organoids may then be implanted into laboratory animals for further study in vivo.  This provides researchers with better tools to investigate normal organ biology or disease states, such as tumor formation.

References:
1. Ogawa, J., Pao, G. M., Shokhirev, M. N., & Verma, I. M. (2018). Glioblastoma model using human cerebral organoids. Cell Reports, 23(4), 1220-1229.
2. Trujillo, C.A., et al. (2019). Complex oscillatory waves emerging from eortical organoids model early human brain network development. Cell Stem Cell, 25(4), 558-569.
3. Sugimoto, S., et al. (2018). Reconstruction of the human colon epithelium in vivo. Cell Stem Cell, 22(2), 171-176.
4. Fujii, M., Matano, M., Nanki, K., & Sato, T. (2015). Efficient genetic engineering of human intestinal organoids using electroporation. Nature Protocols, 10, 1474-1485.
5. Lancaster, M. A., & Knoblich, J. A. (2014). Generation of cerebral organoids from human pluripotent stem cells. Nature Protocols, 9, 2329-2340.

Come visit BTX at the 2020 Society for Neuroscience Annual Meeting, October 24-28, Washington, DC.

Electrical Resistance and Conductance

By Michelle M. Ng, Ph. D.

In this post we will explore the relationship between electrical resistance and electrical conductance. We will also provide information on how resistance applies to electroporation applications, and finally we will answer a FAQ on the topic of sample resistance values in electroporation instrument log data. 

Click here to visit our Electroporation Education Resource, for more basics.

Guidelines for Bacterial Cells

By Michelle M. Ng, Ph. D.

Welcome to the ninth in our series providing tips for developing and improving your own electroporation method—Guidelines for Bacterial Cells.

In this episode, we will touch upon best practice considerations for transforming bacteria including cell preparation, electroporation wave form considerations, cuvette selection, and pulse length.

NOTE: If you have purchased commercially available electrocompetent bacteria, start with the manufacturer's recommended electroporation parameters, and then optimize from there.

NOTE: If you are preparing your own electrocompetent bacteria:

• Grow your cells up to mid-log phase. (For more information, refer to our earlier post Episode 6 - Optimizing Cell Density).
• Wash them well with a low conductance buffer, such as sterile deionized water with 15% glycerol. (For more information, refer to our earlier post Episode 7 - Buffer Considerations).

You may access our protocol for preparing electrocompetent bacteria here.

Tips to boost your success transforming bacteria via electroporation:

Electroporating cloning reactions, such as ligation mixtures, directly into bacteria can cause arcing and reduced transformation efficiencies. 

• To reduce this risk, dilute ligations at least 1:5 with sterile nuclease-free water.
• Alternatively, you can remove salts by dialysis, ethanol precipitation, or spin column clean up.

Bacteria typically work best with exponential decay wave generators. However, you can also electroporate them with a square wave generator. (For more information, refer to our earlier post Episode 1 - Converting Between Square and Exponential Decay Waves).

Most often, 1 mm gap cuvettes are used for bacteria to achieve a high field strength during electroporation. 2 mm gap cuvettes or flatpack chambers may also be used for bacterial electroporation in larger volumes. (Refer to Episode 3 - Scaling Up and Down for more discussion on how the gap sizes affect the field strength). 

Target a field strength range of 15-25 kV/cm (or 1500 to 2500 V using 1 mm gap cuvettes).

You can access our protocol for optimizing bacterial transformation here. This protocol runs through a voltage optimization experiment in detail.

Typically a pulse length of around 5 ms works well for electroporation of bacteria. You may adjust the pulse length by increasing or decreasing the resistance and/or capacitance settings on your exponential decay wave generator.

• T=RC where the T is the Time Constant (time for the voltage to decay to 1/3 the set voltage), R is the Resistance and C is the Capacitance. 

Click here to visit our Protocol Database, for electroporation protocols searchable by, system, cell/tissue type, application/transfectant, and citation.

Guidelines for Mammalian Cells

By Michelle M. Ng, Ph. D.

Welcome to the eighth post in our series providing tips for developing and improving your own electroporation method—Guidelines for Mammalian Cells.

Don’t reinvent the wheel! If there is a protocol or published reference with electroporation parameters for your specific cell line/type (or for a similar cell type), start with it!

You can search for protocols by cell type in our database here.

General Guidelines for Mammalian Cell Electroporation

• Most mammalian cells work best with square wave generators. However, you can also electroporate them to less efficiency with an exponential decay wave generator. (For more information, refer to Episode 1 - Converting Between Square and Exponential Decay Wave).
• Generally 2 mm and 4 mm gap cuvettes are used for mammalian cells. However, 1 mm gap cuvettes may be used for smaller volumes, if the voltage is reduced accordingly, to achieve the desired field strength during electroporation (For more information, refer to Episode 2 - Converting Between Different Gap Sizes).
• Typically field strengths of 400 to 1000 V/cm are used for mammalian cells, and a single pulse ranging from 5 to 25 ms is used for DNA electroporation.
• If using a 2 mm gap cuvette, the voltage ranges from 120 to 200 volts.
• If using a 4 mm gap cuvette, the voltage ranges from 170 to 300 volts.
• For small molecules, such as siRNA, high voltage microsecond pulse lengths may be needed.
• Mammalian cells are best for transfection with a low passage number (ideally passage 3 to 30) and actively growing. Using similar passage numbers between experiments can help to ensure reproducibility.
• Refer to Episode 6 - Optimizing Cell Density for some discussion on optimal cell densities for growth prior to electroporation and suspension at the time of electroporation.
• Refer to Episode 7 - Buffer Considerations for information on using an electroporation buffer such as BTXpress Cytoporation Medium T to boost electroporation efficiency and cell viability.
• High quality DNA that is low in endotoxin and concentrated to 1 to 5 mg/ml is ideal. Prepping DNA with a midi- to maxi-prep, using silica column or anion exchange column technology, is best. The DNA should be dissolved in nuclease-free water as Tris/EDTA solutions can reduce electroporation efficiency.
• It also helps to determine the optimal time after electroporation to analyze your cells.
• For siRNA/miRNA knockdowns, typically 48 hours after transfection is the peak of mRNA knockdown. However, depending on the half-life of the protein, its peak knockdown may occur after that.
• Peak expression of protein from plasmid DNA or mRNA transcript usually falls within 12 to 72 hours after transfection. It is best to run a time-course experiment for each new construct to determine peak expression time.

Please stop by our Blog again soon. Next, in Episode 9, we will explore Protocol Ideas for Bacteria.

Click here to visit our Protocol Database, for electroporation protocols searchable by, system, cell/tissue type, application/transfectant, and citation.

Buffer Considerations

By Michelle M. Ng, Ph. D.

Welcome to the seventh post in our series providing tips for developing and improving your own electroporation method—Buffer Considerations.

The buffer to suspend cells and transfectant at the time of transfection can markedly affect viability of the cells or the stability of the transfectant.  Here are some buffer recommendations based on cell type.

For Bacterial Cells, a low conductance, high resistance buffer is necessary to prevent arcing at the high voltages required to electroporate these cells.

Sterile, distilled water with 10% glycerol is commonly used for bacteria. The glycerol also serves as a cryoprotectant if you plan on freezing cells for future use.

Important tip: When preparing your own electrocompetent cells, be sure to do enough cell washes to remove all residual salts from the growth medium.

You may access our electrocompetent bacteria preparation protocol here.

For Mammalian Cells, specialized, low conductance electroporation buffers are recommended to minimize sample heating and to improve cell viability.

This type of buffer is especially helpful in high voltage applications or when scaling up to larger electroporation volumes. (For more information, refer to Episode 3 - Scaling Up and Down.) An example of a low conductance electroporation buffer is BTXpress Cytoporation Medium T.

For low voltage, cuvette-based applications, often times culture medium (for example RPMI, Opti-MEM) or buffered salt solutions (such as PBS or HBSS) are used. However, since culture media and buffered salts have high electrical conductance, they may cause arcing or undesirable heating.

Important tip: It is critical to leave antibiotics out of electroporation media, to avoid toxicity to cells.

Stay tuned for Episode 8, in which we will tackle Protocol Ideas for Mammalian Cells.

Click here to visit our Protocol Database, for electroporation protocols searchable by, system, cell/tissue type, application/transfectant, and citation.

Optimizing Cell Density

By Michelle M. Ng, Ph. D.

Welcome to the sixth post in our series providing tips for developing and improving your electroporation method—Optimizing Cell Density.

Cell density can significantly affect transfection efficiency. Specifically, if the cells are too dilute or too dense at the time of harvest or at the time of transfection, then viability or efficiency may be compromised. Here are some guidelines to help achieve optimal cell densities by cell type.

Cell density recommendations for harvest:

• Mammalian cells should be grown to a density of 70 to 80% confluency for adherent cells, or 1 to 2 million cells per ml for suspension cells, prior to harvesting for electroporation.
• Bacteria should be grown up to mid-log phase (around OD₆₀₀=0.5) prior to preparation of competent cells.

Cell density recommendations for electroporation:

• For mammalian cells, 5 to 10 million cells per ml is an optimal cell density range for electroporation of most mammalian cells.
• Self-prepared bacterial competent cells are typically in the density range of 1 X 10¹¹ to 1 X 10¹² cells per ml.

Consider running a titration experiment to determine the optimal number of cells to use. To do this, simply test a range of cell densities while keeping the transfectant amount, buffer type, and pulse generator settings the same. At the completion of the experiment, assess the transfection efficiency and cell viability for each cell density.

Stay tuned for our next post, Episode 7 - Buffer Considerations

Click here to visit our Protocol Database, for electroporation protocols searchable by, system, cell/tissue type, application/transfectant, and citation.

Considering Transfectant Amount

By Michelle M. Ng, Ph. D.

This is the fifth post in a series where we are providing tips for developing or improving your own electroporation method.

Transfectant is the molecule of interest that you wish to deliver to your target cells or tissues. The amount of transfectant used in your experiment can impact transfection efficiency. Increasing the amount of transfectant can boost efficiency up to a point; determining the optimal amount to use by running a titration experiment can be helpful.  To do this, you would want to test a range of transfectant amounts while keeping cell number, buffer, and pulse generator settings the same.  Then at the completion of this titration experiment, the transfection efficiency and cell viability for each transfectant amount may be assessed.

Note: The optimal transfectant amount may vary from one construct to another, so it is always a good idea to run a titration experiment each time you begin transfecting a new nucleic acid, protein, or other molecule of interest.

Typical ranges of transfectant for mammalian cells are as follows:

• DNA: 5 to 20 µg/ml (although sometimes increasing to high concentrations such as 50 to 100 µg/ml may enhance transfection efficiency)
• siRNA or miRNA: 5 to 100 nM final concentration. The optimal concentration will be a balance of achieving knockdown vs. minimizing off-target effects. Depending on the potency of the particular siRNA or miRNA and the types of stabilizing modifications on the molecule, more or less may be needed. Modified siRNA typically are used at a final concentration in the 5 to 20 nM range whereas unmodified are more commonly used at a final concentration in the 50 to 100 nM range.
• mRNA 1 to 10 µg per million cells
• Protein 1 to 10 µg per million cells

Tip: Dilute your transfectant in sterile nuclease free water to prevent arcing due to salts or reduced efficiency caused by EDTA or Tris containing buffers.

Coming next is Episode 6 - Optimizing Cell Density

Click here to visit our Protocol Database, for electroporation protocols searchable by, system, cell/tissue type, application/transfectant, and citation.

 Design Your Own Electroporation Protocol Episode 4 - 

Temperature Considerations

By Michelle M. Ng, Ph. D.

This is the fourth in a series of posts where we will be providing tips for developing or improving your own electroporation method.

The majority of mammalian cells are electroporated efficiently at room temperature.  However there are couple reasons to consider an electroporation experiment at a lower temperature such as 4  ͦC:

• To reduce cell heating. As mentioned earlier in Episode 3 as you increase volume you may also increase conductance and heating of the sample which can have negative effects on cell viability. Electroporation programs with high voltage pulses (typically with bacteria), long pulse durations or multiple pulses may also cause heating of the cells

• To keep pores open longer after electroporation. Since electroporation causes the transient formation of pores, keeping the cells at a lower temperature following the pulse may allow the pores to remain open longer to allow more uptake of the transfectant.

One option is to include pre-chill and post-chill steps by placing the cells/cuvettes on ice or in the refrigerator for a few minutes before and after electroporation.

• Make sure you wipe off any condensation on the outside of the cuvettes before placing it in the safety stand, safety dome or PEP.
• The standard electroporation pulse voltage used for cells at room temperature will need to be approximately doubled for electroporation at 4 ͦC to effectively permeate the cell membrane.

Example: If you typically electroporate cells at 250 V at room temperature, then you would want to electroporate the same cells at 500 V at 4 ͦC.

Note: temperature can have big effects on outcome of your experiments. If you are not certain which temperature will work best for your cells it might be a good idea to test a few replicates of different temperature conditions in a single experiment such as room temperature, chilled to 4 ͦC, and chilled on ice side by side with the same electroporation parameters, cells, and transfectant kept constant.

Come back next time when we look at Episode 5 - Considering Transfectant Amount

Click here to visit our Protocol Database, for electroporation protocols searchable by, system, cell/tissue type, application/transfectant, and citation.

By Michelle M. Ng, Ph. D.

This is the third post in a series where we are providing tips for developing or improving your own electroporation method. 

There is some room for scaling electroporation volumes up and down just by switching cuvette gap sizes.

• 1 mm gap cuvettes accommodate small volumes (20 to 90 µl)
• 2 mm gap cuvettes accommodate medium volumes (40 to 400 µl)
• 4 mm gap cuvettes accommodate larger volumes (80 to 800 µl)

The process of scaling up and down between different sized chambers of roughly the same dimensions (such as cuvettes) is simple. You just want to adjust your voltage while keeping the field strength in V/cm and other parameters constant.

Example: You typically electroporate your cells for a single pulse, 5 msec, 250 V in a 2 mm gap cuvette, but would like to convert to a 1 mm or 4 mm gap cuvette. When converting these settings from a 2 mm gap to a 4 mm gap cuvette, you would want to electroporate your cells for a single pulse, 5 msec, 500 V. And in order to convert these settings from a 2 mm gap cuvette to a 1 mm gap cuvette, you would want to electroporate your cells for a single pulse, 5 msec, 125 V.

For even larger volumes you will need to move to a different type of chamber. You may use the same rules above about adjusting voltage parameters to achieve the same field strength when moving from cuvettes to a different style chamber, however keep in mind you may also need to do some protocol fine tuning to optimize for the different geometry of the chamber. For reference, the parameters normally used for a 4mm gap cuvette would also be a good starting point for a 10 ml volume, 4 mm gap large volume electroporation flatpack.

Note: Keep in mind as you increase volume you also increase conductance of your sample. This means more heating of the cells during the electroporation pulses which can cause cell viability issues. To combat this effect you may want to consider using a low conductance electroporation buffer and/or experiment with prechilled buffer for larger volume electroporations.

Our next post in this series will be Episode 4 - Temperature Considerations

Click here to visit our Protocol Database, for electroporation protocols searchable by, system, cell/tissue type, application/transfectant, and citation.

Visit BTX at Immunology 2019, May 10-12, San Diego, CA, Booth #1417

This is the second in a series of posts where we are providing tips for developing or improving your own electroporation method. 

Today's post will be short and sweet, as we explore how to convert between two electrodes or cuvettes with different gap sizes.

Gap distance is defined as the distance between electrode contacts or between parallel electrode plates in a cuvette.

Electrical Field strength (expressed as Volts/centimeter or kiloVolts/centimeter) is equal to the voltage (V) set on your electroporator divided by the gap distance between your electrode contacts (cm).

This means that the relationship between gap distance and field strength is inversely proportional; as gap distance increases, electrical field strength decreases.

The simple answer to convert an electroporation protocol for different electrode gap distances is to adjust the voltage such that the field strength (in V/cm) is constant between two different gap sizes, while keeping the other parameters such as pulse length and number of pulses the same.

As an example, if you normally electroporate at 500 V in a 4 mm gap cuvette, then you would want to use 250 V for a 2 mm gap cuvette or 125 V for a 1 mm cuvette; all of these would achieve a final field strength of the same 1250 V/cm.

Note: If the electrode is a different shape or there is a change in the relative volume of your sample, some additional fine tuning may be necessary for best results.

Next time we'll tackle Episode 3 - Scaling Up and Down
 

Click here to visit our Protocol Database, for electroporation protocols searchable by, system, cell/tissue type, application/transfectant, and citation.

In the next series of posts we will be giving tips for developing or improving your own custom electroporation method. The topics we will be covering in this series are:

Episode 1- Converting Between Square and Exponential Decay Waves

Episode 2- Converting Between Different Electrode/Cuvette Gap Sizes

Episode 3- Scaling Up and Down

Episode 4-Temperature Considerations

Episode 5-Transfectant Quantity

Episode 6- Cell Density

Episode 7- Buffer Considerations

Episode 8- General Protocol Ideas for Mammalian Cells

Episode 9- General Protocol Ideas for Bacterial Cells

Today we will explore Episode 1- Converting Between Square and Exponential Decay Waves.

Episode 1 - Converting Between Square and Exponential Decay Waves

Note: Please see our earlier post, Catching the Electroporation Wave, for an explanation of square and exponential decay waveforms.

Converting from Exponential Decay Wave (ECM 399, ECM 630, Gemini) to Square Wave (ECM 830, Legacy ECM 2001, ECM 2001+): Use the same voltage and select a square wave pulse length of 1/2 the exponential decay wave time constant.

1. Select the appropriate square wave electroporation mode on the instrument.
2. Keep the square wave voltage, gap size, pulse number, and pulse interval parameters the same as the exponential decay wave protocol.
3. No need to set Resistance (R, Ω) or Capacitance (C, mF) on square wave instruments
4. Set the square wave pulse length to one half the time constant of the exponential decay wave protocol.

Converting from Square Wave (ECM 830, Legacy ECM 2001, ECM 2001+) to Exponential Decay Wave (ECM 630, Gemini): Use the same voltage and adjust the exponential decay wave capacitance and resistance settings to achieve a time constant of 2X the square wave pulse length.

1. Select the appropriate exponential decay wave electroporation mode on the instrument.
2. Keep the exponential decay wave voltage and gap size parameters the same as the square wave electroporation protocol.
3. Set the pulse number: 1 to 2 pulses are available for single sample mode on ECM 630 and Gemini 7" touchscreen instruments.  For Legacy ECM 630 instruments, only a single pulse is available per experimental protocol.  For protocols requiring more than 1 to 2 pulses, additional pulses may be executed by manually running the protocol more than once sequentially.
4. Adjust the instrument resistance (R, Ω) and Capacitance (C, μF) settings on the exponential decay wave generator to achieve a time constant that is double the square wave pulse length

These conversions are theoretical as generally the square wave works best for mammalian cells and exponential decay waves work best for bacteria and yeast.

Please note you may need to do some additional optimization and be willing to accept some drop in performance when using a different waveform than is optimal for your cell type.

For an additional explanation of this theoretical conversion between the two waveforms and some universal electroporation protocols for mammalian cells in both square wave and exponential decay wave, please read the document attached to this link: http://www.btxonline.com/media/wysiwyg/protocol_db/Theoretical_Conversion.pdf

Our next episode is Episode 2 - Convert Between Different Electrode/Cuvette Gap Sizes

Click here to visit our Protocol Database, for electroporation protocols searchable by, system, cell/tissue type, application/transfectant, and citation.

Improving Cell Viability During Transfection 

 By Michelle M. Ng, Ph. D.

There are several common causes of cellular toxicity following electroporation.  Here’s a list of the top causes of cell death during electroporation experiments and recommended solutions.

• Sub-optimal electroporation buffer. Buffers with a high salt content such as PBS, Opti-MEM, and RPMI are also highly conductive and can cause overheating and cell death during electroporation.  Switching to a specialized low conductivity electroporation buffer such as Cytoporation Medium T can improve cell viability following electroporation.
• Cells not transferred immediately to culture vessel after electroporation. If you are electroporating in a buffer other than the complete media used to culture your cells, it is best to transfer the cells from the cuvette to a culture dish containing growth medium immediately after each electroporation. This will minimize stress to the cells.
• Electroporation pulse voltage is too high. Increasing voltage can boost electroporation efficiency up to a point, however increasing voltage can also increase cell toxicity. To improve viability, decrease the voltage by increments of 10 volts to improve viability.  Oftentimes, it is helpful to run an optimization experiment at a range of different voltages and assess electroporation efficiency and viability at each.
• Electroporation pulse length is too long. Although increasing pulse length can boost electroporation efficiency in some cases, increasing pulse length can also increase cell death. Decrease the voltage and pulse length by increments of 10 volts and 2 to 5 ms when using a square wave generator. Decrease the voltage and capacitance by increments of 10 volts and 100 µF when using an exponential decay generator.
• Cell density too low or too high. Typically a density of 1 to 10 million cells per ml is optimal at the time of electroporation.  However it is always a good idea to test several different cell densities at the time of electroporation for your particular cell type to determine what works best. Optimizing the number of cells to plate for your culture vessel size following electroporation can also improve viability.
• Passage number too low or too high. It’s best to allow freshly thawed cells to recover and go through 2 to 3 passages before using them for transfections. Also it is best to avoid high passage cells (> passage 30).
• Contaminated transfectant. Use highly purified, sterile, contaminant-free DNA or RNA with low endotoxin levels. Contaminating salts, organics (such as ethanol carry over from precipitations and spin columns), endotoxin, or pathogenic microorganisms can contribute to viability issues. Also, it is critical to exclude antibiotics from the electroporation medium and allow adequate recovery time before re-adding selective media post-electroporation.

Click here to visit our Electroporation Education Resource, for applications information, guides to technology selection and protocol optimization, references, and more

Unleash the Potential of CRISPR with Electroporation 

 By Michelle M. Ng, Ph. D.

The CRISPR/Cas system is a prokaryotic immune system that allows the cell to protect itself from foreign DNA such as a virus or plasmid. Modified versions of CRISPR/Cas9 are being utilized as gene editing tools in revolutionary ways in science and medicine.

 Part of this system has been modified to be used as a tool for editing genomes.  A small guide RNA (sgRNA, 1) is employed which is complimentary to a PAM recognition site in the target area of the genome.  Next the sgRNA and Cas9 protein form an ribonucleoprotein complex (2, RNP) which is capable of causing sequence-specific double stranded breaks at the PAM site (3).

These double stranded breaks can either be faithfully repaired or induce point mutations by DNA repair mechanisms of the cell (4, leading to frameshift mutation/knock-out) or be used to replace with DNA of interest at that site (Homology directed repair, HDR). 

Here we explore the formats that CRISPR/Cas9 tools and the benefits of using electroporation to deliver CRISPR gene editing constructs.

What Forms of CRISPR Constructs? And Why?

• Plasmid based CAS9 and gRNA expressed from one construct. For this the expression promoters for the Cas9 protein and gRNA have to be compatible with your cell type.  The plasmid lasts longest in the cell so the potential for off target effects are the greatest with this format. Because both the Cas9 protein and the gRNA have to be made and find each other in the cell to produce a functional RNP, this is also the least efficient format.
• mRNA , CAS9 and gRNA supplied in trans. This format is not cell type specific. mRNA format has an intermediate level of efficiency and has an intermediate half-life in the cell. Also the mRNA format has less chance of off target effects than plasmid format. Working with the mRNA can be daunting to some because RNA is sensitive to degradation.
• Protein CAS9 and gRNA precomplexed and delivered as an RNP. This is the most efficient method because the RNP is already ready to act once delivered to the cell and also is around for the shortest period of time (hours).  Protein format also has the least risk of off-target effects
• Additional molecules for knock-ins: ssODN or dsDNA (PCR products or plasmid) for HDR template are added in addition to the CRISPR formats listed above. Short single stranded oligos (ssODNs) may be used for small changes such as a single amino acid substitution, however plasmids are most commonly used as HDR templates for larger knock-ins such as whole genes or florescent reporters.

Why Electroporation is an ideal delivery method for CRISPR

• Electroporation is an efficient method of delivering large or complex mixtures of molecules that were previously impossible to transfect by traditional chemical, lipid, and viral delivery methods.  Multiplexed CRISPR gene editing and whole gene knock-ins are within reach with this technology.
• Electroporation is a physical method of transfection, and works well even with difficult to transfect samples, such as primary cells, suspension cells, embryonic cells, or even whole tissues in vivo.  For 3D cultured cells or slices of tissue ex-vivo, the sample may be placed into a specialized electrode chamber.  For in vivo transfection, the target tissue of the animal may be injected with the transfectant, then contacted with paddle shaped electrode probes, or reached by electrode needles inserted into the tissue.
• Electroporation pulse parameters are reproducible from one experiment to the next, and are not subject to lot to lot reagent variation or toxicity issues. The electrical pulses themselves are easy to monitor during the experiment using either an advanced electroporation instrument with pulse monitoring and logging features or attaching a high voltage probe and oscilloscope. Following electroporation, transient cell pores seal up leaving behind no residual chemicals that may affect cell viability.

Click here to learn more and download our latest CRISPR application note

The Immune system is the body’s defense against infections and cancer. It consists of billions of cells that are divided into several different types. A subtype of white blood cells calls Lymphocytes comprise a major portion of the immune system. There are three types: B lymphocytes, T lymphocytes and Natural Killer (NK). In today’s blog post we focus on T lymphocytes (T cells) and how they work to kill tumors. 

CAR-T stands for Chimeric Antigen Receptor T-Cells which are custom made T cells used to attack your target of choice. One common target is specific receptors on tumor cells. Currently a favored method is to transfect mRNA into T-cells using electroporation. 

The difficulty associated with non-viral gene transfer methods in primary lymphocytes can result in low gene transfer efficiency and high transfection related toxicity. Improved transfection efficiency was achieved by electroporation using in vitro transcribed mRNA. RNA electroporation may be used to engineer T cells with new biological functions, providing a new and powerful tool for altering T cell biology where long term transgene expression is not necessary or desirable. 

RNA Electroporation was found to improve transfection efficiency (90% achieved with RNA as compared with about ~50% efficiency for plasmid DNA; 1,2) and reduced transfection related toxicity. Optimization experiments using different electroporation parameters for stimulated human PBLS  that viability of transfected T cells was 63-86% range 24 hours after electroporation. Cells stimulated from 2 to 18 days showed similar efficiencies indicating that the post stimulation time length does not greatly influence RNA electroporation. RNA electroporation shows advantages over plasmid DNA based electroporation. Optimized electroporation conditions did not induce adverse effects on T-cells viability and proliferation or cause apoptosis. 

In conclusion, RNA electroporation is highly efficient tool to introduce genes into human and murine primary T lymphocytes. This technology can be used to engineer T cells with new biological functions. 

Stay tuned as next time we tackle a 9 part series on “Designing your own optimized Electroporation Protocol”

1. Zhao Y, Zheng Z, Cohen CJ, Gattinoni L, Palmer DC, Restifo NP, Rosenberg SA, 
   Morgan RA. High-efficiency transfection of primary human and mouse T lymphocytes 
   using RNA electroporation. Mol Ther. 2006 Jan;13(1):151-9. Epub 2005 Sep 2. 
   PubMed PMID: 16140584; PubMed Central PMCID: PMC1473967. https://www.ncbi.nlm.nih.gov/pubmed/16140584
2. Bell MP, Huntoon CJ, Graham D, McKean DJ. The analysis of costimulatory 
   receptor signaling cascades in normal T lymphocytes using in vitro gene transfer 
   and reporter gene analysis. Nat Med. 2001 Oct;7(10):1155-8. PubMed PMID: 
   11590441 https://www.ncbi.nlm.nih.gov/pubmed/11590441

Exponential Decay versus Square

By Michelle M. Ng, Ph. D.

Electroporators can come in multiple electrical waveform styles and typically allow you to vary the characteristics of the electrical pulse settings. Every cell type is unique in terms of which pulse characteristics work best. Field Strength (€, usually expressed in V/cm) is dependent on the pulse parameters applied (voltage, capacitance and resistance) and the distance between the electrode or cuvette contacts. Application of this electrical field causes Electropermeabilization (transient pores in the cell membrane through induction of transmembrane voltage) allowing nucleic acids to pass through the cell membrane.

The two main different types of pulses used for electroporation of nucleic acids:

• Exponential Decay Wave is typically used for cells with cell walls such as bacteria and yeast. The generator hits the peak voltage at the beginning of the pulse then decreases over time.

• Square Wave is generally used for mammalian cells and tissues at a lower field strength than exponential decay. The generator jumps up to the set voltage and holds steady for the desired pulse length. 

Whether you are looking to ride a short or long wave make sure you make note of dependents that factor in to your labs work. Check back next time when we “Attack the Target using CAR-T cells. 

Transfection is simply getting DNA (or other molecules of interest) into a cell, while keeping that cell alive. We can do this in countless methods which are divided into three main categories; Reagent Based, Instrument Based, Biological Based. Each method has its own advantages and disadvantages and may be used for transient or stable transfections. Read on as we indulge ourselves into the top methods to see which one suits you best. 

CA Phosphate
This reagent based method involves mixing DNA with calcium chloride, adding in a precise manner to a buffered saline/phosphate solution and allowing the mixture to sit at room temperature. This produces a precipitate that is isolated onto the cultured cells via endocytosis or phagocytosis.

Advantages:

• Inexpensive
• High efficiency in compatible cell types

Disadvantages:

• Reagent uniformity is critical for reproducibility; difficult to achieve reproducibility from one experiment to the next
• Small pH changes (+0.1) can compromise alteration efficiency
• Size and quality of the precipitate are vital to the success of transfection
• Does not work well for all cell types
• Toxicity due to lingering reagent after transfection
• Calcium phosphate precipitation does not work in RPMI, due to the high focus of phosphate within the medium.
• Efficiency is limited by the size of DNA molecules

Lipid Based Methods
This reagent based method helps a cell to absorb nucleic acid by neutralizing its negative charge and making the molecule more compatible to getting past the hydrophobic cell membrane (1). Genetic material is transferred into the cell via liposomes; these are vesicles that can merge with the cell membrane.

Advantages:

• Proficiency to deliver nucleic acid to many adherent cell lines
• Low toxicity in many easy to transfect cell lines-deliver with low or little cell death
• Easy to use- minimal steps
• Low start-up costs

Disadvantages:

• Not effective with many difficult-to-transfect, suspension, and primary cell types, and not applicable to cells with cell walls such as bacteria, yeast, and plant cells.
• Toxicity due to lingering reagent after transfection in sensitive cell lines
• High consumable costs for experiments over time
• Lot to lot reagent variability in results
• Reagent is sensitive to storage conditions
• Efficiency is limited by the size of DNA molecules

Viral Vectors
This biological based method uses Viral vectors (such as retrovirus, lentivirus, adenovirus or adeno-associated viruses) to deliver nucleic acid into the cells.

Advantages:

• High gene-delivery efficiency (95-100%)
• Simplicity of infection

Disadvantages:

• Labor intensive preparation of virus
• Virus is sensitive to storage conditions
• Some viruses (such as lentivirus) can cause unwanted mutations to the host cell’s genome
• Increased Biosafety Concerns--P2 containment required for most viruses (certain viruses can be dangerous and should be closely monitored.)
• Viral packaging limits prevent delivery of large constructs 

Electroporation
This instrument based method uses an electrical pulse to create brief pores in cells membrane through which substances like nucleic acids can pass into cells (2).

Advantages:

• Non-chemical method that doesn’t seem to alter the biologic structure or function of the target cells; no lingering reagent after transfection can improve viability in reagent sensitive cell types.
• Easy to perform
• High Efficiency, High Viability
• High Reproducibility of results from experiment to experiment
• Universal transfection method that can be applied to a wide range of cell types in vitro, as well as in vivo tissue transfections, and can be used to deliver a variety of different transfectant cargo molecules or nanoparticles
• Consumable cost savings over time
• Efficiency of this method is not limited by size of DNA molecules

Disadvantages:

• Cell mortality (if using suboptimal conditions)
• Higher start-up costs

Gene Gun
This instrument based transfection method uses nitrogen or helium gas for injecting tiny ‘bullets’ made from DNA coated particles, mainly used on plants and for in vivo animal applications.

Advantages:

• Consumable Cost savings over time
• Transfectant applied with or without gold particles.

Disadvantages:

• Not applicable to in vitro cells
• Causes damage in target tissue
• Trial animals easily shocked by the sonic boom
• High start-up costs
• Labor intensive multi-part procedure in preparing samples

Microinjection
This instrument based method uses a fine needle to deliver nucleic acid into the cytoplasm or nucleus one cell at a time.

Advantages:

• Cell type independent
• Uses smaller amounts of DNA
• Efficiency of this method is not limited by size of DNA molecules
• High reproducibility

Disadvantages:

• Labor intensive, limited to small numbers of cells, and requires highly skilled laboratory personnel
• High Instrument costs
• Causes physical damage and high mortality to the samples

Mission Impossible may not be so impossible after all. These methods should help you narrow down how you wish to proceed in your labs. Check back next week when we try to “Catch the Electroporation Wave”.  

References:
1: Kepczynski, M., & Róg, T. (2016). Functionalized lipids and surfactants for specific applications. Biochimica et Biophysica Acta (BBA)-Biomembranes, 1858(10), 2362-2379

2: Rosazza, C., Haberl Meglic, S., Zumbusch, A., Rols, M. P., & Miklavcic, D. (2016). Gene electrotransfer: a mechanistic perspective. Current gene therapy, 16(2), 98-129