Automated IDT Alt-R CRISPR/Cas9 Ribonucleoprotein Lipofection Using the Biomek i7 Hybrid Automated Workstation

Abstract

The ability to precisely edit the genomes of host species using CRISPR/Cas9 has revolutionized how the scientific community performs genetic manipulations, and CRISPR screening has become a popular method to evaluate how changes in genotype can affect a phenotypic outcome. In order to make precise edits in the genome, CRISPR systems use a ribonucleoprotein (RNP), which contains a Cas9 enzyme bound to a guide RNA (gRNA). The Cas9 nuclease introduces DNA breaks at regions defined by the gRNA. There are numerous ways to introduce RNPs into the cells of interest, but one method that has become standard within the field is lipofection, which is the delivery of genetic material using liposomal transfection. During CRISPR lipofection, recombinant Cas9 protein is combined with a gRNA in a cationic lipid nanoparticle to generate functional RNPs, which are then introduced into living cells. Here we sought to automate the delivery of IDT Alt-R CRISPR/Cas9 RNPs into HEK293 cells using lipofection. We compare our newly developed method on a Biomek i7 Hybrid Automated Workstation method with results obtained by manual, hands-on experiments. Using fluorescently labeled gRNA, we show that transfection efficiency is comparable between the two methods, but that the gentle, reproducible pipetting performed by automation provides more precise results. In addition, using the IDT Alt-R Control Kit, we show the automated method can edit genes with an efficiency like manual experiments. Together this data demonstrates an automated method for lipofection of IDT Alt-R RNP into HEK293 cells and the analysis of the resulting mutations using the IDT T7EI system. The automated method performs comparably to manual experiments, with similar editing efficiencies, while requiring less hands-on time and reducing the chance of user-introduced error.

Introduction

Since its Nobel Prize-winning discovery in 2011, CRISPR/Cas9 has become a vitally important research tool within the biomedical community.^1,2 The CRISPR/Cas9 system was adapted from a naturally occurring bacterial immune process and has evolved to allow researchers to make precise genetic edits in a variety of host species.1 Now basic researchers can rapidly make complex genetic models of disease and explore the relationship between genotype and phenotype. Additionally, there is hope that one day this technology can even be extended to the clinic, where CRISPR could be used to treat heritable disorders that occur due to defined point mutations, but further work in this area remains to be done.

The CRISPR/Cas9 RNP is made of two main components: a nuclease (Cas9), which introduces double-stranded DNA breaks, and a guide RNA (gRNA) that directs the Cas9 to the correct genomic locus. The gRNA can be further divided into the tracrRNA, which is responsible for binding of Cas9 to the RNA, and the crRNA, which targets the RNA-Cas9 to the genetic region of interest. The gRNA can be a single piece of RNA, referred to as a single guide RNA (sgRNA), or the tracrRNA and crRNA can exist as separate strands in a two-part gRNA system.3 The double-stranded breaks introduced by Cas9 can be repaired via two mechanisms: non-homologous end-joining (NHEJ) or homology directed repair (HDR) when a repair template is present. NHEJ cause imprecise repairs, yielding insertion/deletions (indels) of varying sizes at the location of the DNA break. The indel patterns are not random and differ from gRNA to gRNA. HDR offers the opportunity for insertion of new sequences at the DNA break site.

These new sequences can be delivered into cells as a donor template, can range in size from 1 to 1000s of bases, and can code for entirely new genes.

There are several ways that functional RNPs can be introduced into cells, such as through engineering of cell lines or viral infection, but the most common ways are via electroporation or lipofection. Electroporation uses an electrical field to make the membranes of cells more porous to allow the introduction of exogenous material, like RNPs. While this option is useful in cell lines that are hard to transfect, it has the drawbacks of being expensive and requiring specialized laboratory equipment. The other option is lipofection, which forms the basis of IDT’s Alt-R CRISPR/Cas9 system. This system combines purified, recombinant Cas9 protein with a synthetic gRNA to form RNPs. This system is relatively fast and straightforward and uses common, commercially available transfection reagents, like Lipofectamine (Invitrogen), to introduce the RNPs into the cell line of interest (Figure 1A).⁴

Briefly, RNPs are assembled by incubating the Cas9 protein with a crRNA: tracrRNA duplex and combining this mixture with commonly used transfection reagents. This mixture is then added to a cell culture plate and used to reverse transfect the cell line of interest. Transfection efficiency and gene editing can then be assessed using fluorescence microscopy (if labeled tracrRNA is used) and IDT’s T7 endonuclease I (T7EI) cleavage assay, respectively (Figure 1B). In the T7EI assay, the genomic region where mutations were introduced is amplified by PCR, and the resulting fragments are denatured and allowed to slowly anneal to form dsDNA. As both wild type and mutated amplicons are generated, some of these dsDNA duplexes will have mismatches in the mutated region (heteroduplexes). The T7EI enzyme will cleave the dsDNA at these mismatches, generating DNA fragments that can be separated and quantified to estimate CRISPR editing efficiency. The T7EI assay tends to underestimate editing efficiency as 1 base pair insertions/deletions are not detected as efficiently. To fully determine editing efficiency, next generation sequencing is recommended. Nonetheless, the T7EI assay is a straightforward and quick method to estimate editing efficiency.

As CRISPR-based gene editing has become more commonplace, there has emerged a growing interest in using it as the basis of screening. CRISPR screens can be largely divided into two methods: arrayed and pooled screening. The most popular method for pooled CRISPR screening is SLICE.5 In a SLICE workflow, a population of cells is infected with a lentiviral library that contains numerous different gRNAs, followed by electroporation of purified Cas9 protein. This workflow is amenable to numerous cell types, including primary cells, but can be labor and resource intensive, as the workflow is long. Further, data deconvolution can be tedious, as cellular separation and NGS steps are required to identify which gRNAs gave rise to the phenotype of interest.

On the other hand, arrayed screening can be much more straightforward, as each well of a microtiter plate contains a unique gRNA(s) targeted against a gene of interest, so NGS deconvolution is unnecessary. This makes data analysis much more straightforward, as the phenotype in each well can be rapidly evaluated using any plate-based detection assay. This method also gives the researcher more control over what genes to screen against, as targeted gRNA arrays can be designed to probe specific pathways or gene families.6 In addition, there are even commercially designed crRNA libraries readily available from IDT. As arrayed CRISPR screening can potentially involve the use of many microtiter plates, differing only in the identity of the gRNA in each well, it is an especially attractive target for laboratory automation. As evidenced here, automation of IDT Alt-R RNP formation and lipofection using a Biomek i7 Hybrid Automated Workstation has several key advantages over performing experiments by hand.

The Biomek i7 Hybrid Automated Workstation is an automated liquid handler that is capable of efficiently performing the complex liquid handling steps of CRISPR RNP workflows (Figure 2). This minimizes the number of required user interactions and increases walkaway time, freeing the operator to attend to other laboratory tasks. The multichannel pod can be equipped with a 96-well head that can accurately pipette 1 to 1200 μL or a 384-well head that is accurate over the range of 0.5 to 60 μL. Additionally, the 8-channel Span-8 pod is accurate from 1 to 1000 μL. This workstation supports 45 deck positions and can be directly fitted with orbital shakers, heating/cooling Peltiers, and tip-washers for plate and sample processing (Figure 2).

Further, depending on user needs, the Biomek i7 Hybrid workstation supports integration with other automated plate handling instruments, such as thermal cyclers, incubators, barcode readers, washers, multimode plate readers, centrifuges, and more. Another important feature is the optional HEPA filter, which creates a more sterile environment, an important factor when handling mammalian cell cultures. Here, we show that automated IDT Alt-R RNP formation and mutation discovery using a Biomek i7 Hybrid workstation provides excellent results that are equivalent to manually processed samples. The automated workflow can reduce hands-on time and the possibility of sample handling errors by the user.

Methods

Cell Culture

Lenti-X 293T cells (Takara) were maintained at 5% CO₂ and 37°C in growth medium, which was composed of DMEM supplemented with 10% FBS and 1% 100X antibiotic/ antimycotic (Gibco). On the day of lipofection, cells were washed with DPBS, harvested with trypsin, pelleted at 300 x g in a Beckman Coulter Allegra X-14R centrifuge, and resuspended in 10:1 OptiMEM to growth medium. Cells were counted and further diluted to achieve desired cell density of 400 cells/μL.

RNP Formation and Lipofection

Prior to RNP formation, 100 μM stock solutions of human Hprt-targeted and negative control crRNAs and ATTO-550 tracrRNA (IDT) were made using nuclease-free IDTE buffer. Each crRNA was then combined in a 1:1 molar ratio with the tracrRNA and diluted to 1 μM in nuclease-free duplex buffer. Samples were heated to 95°C for 5 minutes and slowly cooled to ambient temperature to generate RNA duplexes. While the RNA complexes were cooling, Cas9 V3 nuclease (IDT) was diluted 62-fold in OptiMEM to achieve a final concentration of 1 μM. RNP formation was then performed manually using handheld pipettes and via an automated method on the Span-8 pod of a Biomek i7 Hybrid Automated Workstation. RNPs for each crRNA were formed by combining 100 μL of 1 μM complexed RNA with 100 μL of 1 μM Cas9 V3 nuclease, 40 μL Cas9 PLUS reagent (Invitrogen), and 1425 μL OptiMEM, and incubating at ambient temperature for 5 minutes.

To assemble lipofection reactions, 23.8 μL OptiMEM and 1.2 μL CRISPRMAX transfection reagent (Invitrogen) were added to each well of a 96-well polystyrene, tissue culture treated plate, either by hand or the automated method. Next, to generate lipofection complexes, the RNPs formed above were added to each well to achieve the desired final concentration of RNP: 0 μL (Negative control), 12.5 μL (0.5X), or 25 μL (1X). OptiMEM was then added so that all wells contained a total volume of 50 μL. Lipofection complex formation was carried out for 30 minutes at ambient temperature. After this incubation, 100 μL of the HEK cell suspension prepared above were added by hand or via the automated method with the Span-8 pod (Figure 3). Plates were incubated at 5% CO₂ and 37°C for 48 hours.

Media Exchange and Imaging

Prior to cell imaging and ATTO 550 fluorescence measurement, cell media was aspirated, either manually or using the Biomek i7 Hybrid workstation, and replaced with 100 μL of DPBS (Gibco). For the automated method, these pipetting steps were performed at a rate of 2 μL/second in order to keep adherent cells attached (Figure 3). Total fluorescence in each well was then measured using a SpectraMax i3X (Molecular Devices) at an excitation wavelength of 550 nm and an emission wavelength of 570 nm. Representative wells of each condition were then imaged using an EVOS-FL system (Advanced Microscopy Group) using both bright-field and an RFP filter cube. All wells were imaged using identical acquisition parameters, including 40% fluorescence intensity and 100 ms acquisition time in the RFP channel.

gDNA Isolation and PCR

To isolate gDNA for mutation analysis, wells containing HEK293 cells were washed with 100 μL DPBS either manually or using the automated method as above (Figure 3). DPBS was then removed and replaced with 50 μL of Quick Extract DNA solution (Lumigen), the plates were shaken at 300 rpm for 3 minutes on the Biomek’s on-deck orbital shaker, and the well contents for both manual and automated plates were transferred to a single 96-well PCR plate (BioRad). Samples were then heated to 65°C for 10 minutes then 95°C for 5 minutes on the Biomek’s Peltiers. All samples were then diluted with 100 μL nuclease-free water and stored at -20°C until the day of the PCR run.

To amplify the human Hprt locus, PCR was performed using the primers supplied in the IDT Alt-R Human Control Kit and KAPA HiFi Hot Start Ready Mix. For each sample, PCR mix was assembled in a BioRad 96-well PCR plate using the Biomek i7 Hybrid workstation by combining 12.5 μL of KAPA Ready Mix with 6.5 μL of 1.15 μM Primer Mix and 6 μL of gDNA harvested above (Figure 3). For all samples, PCR was performed on the Biomek’s on-deck ATC using a protocol with an initial melt at 95°C for 5 minutes, followed by 32 cycles of 98°C for 30 seconds (melt), 67°C for 20 seconds (anneal), and 72°C for 45 seconds (extend). This was followed by a final extension of 2 minutes at 72°C. The plate was then stored at -20°C until the day of the T7EI assay.

T7EI Assay

In order to assess gene editing, a T7EI assay was performed on the PCR amplified Hprt locus. Using the Biomek i7 Hybrid Automated Workstation, PCR products of replicate wells of each condition were combined, and 10 μL of each pooled condition were added to 8 μL of 2.5X T7EI Reaction Buffer (IDT). After PCR product dilution, heteroduplexes were formed using the on-deck ATC. To generate heteroduplexes, the mixtures were heated to 95°C for 10 minutes to denature the DNA. This was followed by cooling from 95°C to 85°C at a ramp rate of -0.4°C/second, then samples were slowly cooled from 85°C to 25°C at a ramp rate of -0.25°C/second.

Next, 2 μL of T7EI enzyme was added to each well using the Biomek’s Span-8 pod, and the reaction was incubated for 1 hour at 37°C in the Biomek’s on-deck ATC (Figure 3). In order to analyze heteroduplex mismatch restriction, T7EI reactions were diluted 1:50 in 0.1X IDTE buffer (IDT) and analyzed using a Bioanalyzer 2100 with the High Sensitivity DNA Kit (Agilent). Editing efficiency was calculated as previously described, using Bioanalyzer calculated molar concentrations and the equation below, where F₁ and F₂ refer to 250 and 850 bp cut DNA fragments, respectively, and FL refers to full-length Hprt amplicon.³ For some samples, several peaks occurred between 220 and 280 bp. In these cases, the concentrations of all peaks within this range were added together to generate a cumulative F₁ value.

Results and Discussion

Automated RNP Transfection

An important step in any CRISPR workflow is the successful delivery of Cas9 protein and gRNA in the form of an RNP into the target cells. Here the Alt-R CRISPR/Cas9 system was used, where a recombinant, purified Cas9 variant was combined with synthetic tracr- and crRNAs. Two crRNAs were used throughout the work described here, both of which were included in the IDT Human Control Kit. The first crRNA was targeted against the human hypoxanthine phosphoribosyltransferase I (Hprt) gene, while the other was a negative control crRNA. The negative control crRNA is designed by IDT and contains a protospacer region that does not target the gRNA for editing in human or murine genomes. Thus, the negative control crRNA should not produce gene edits in the Hprt locus.

The tracrRNA used for these experiments was fluorescently labeled with ATTO 550, which allowed the visualization RNP delivery following transfection (Figure 4). RNPs were formed according to IDT directions via the automated method and by hand, then added to HEK293 cells in 96-well plates. Control wells contained no RNP. Two days after RNP transfection, wells were imaged to evaluate transfection efficiency. Representative images are presented in Figure 4.

Analysis of bright-field images showed that both automated and manually prepared wells contained viable cells with the expected morphology (Figure 4A,B,E,F). Additionally, the wells appeared to contain approximately the same number of cells. Using the same well locations, ATTO 550 fluorescence was assessed in the RFP channel. As shown in Figure 4D and 4H, robust fluorescence was observed in wells containing ATTO 550-labeled RNPs. Conversely, control wells with no RNP exhibited no signal under identical image acquisition settings (Figure 4C and 4G). Qualitatively, there appeared to be no differences observed between assay plates assembled manually and using the Biomek i7 Hybrid automated method (Figure 4). Together this indicated that HEK293 cells could be efficiently reverse transfected using IDT Alt-R reagents on a Biomek i7 Hybrid workstation.

To quantitively analyze well-to-well variability of the transfection methods, ATTO 550 fluorescence in each well was measured using a Molecular Devices SpectraMax i3x plate reader. Two concentrations of RNP were delivered to cells: the amount recommended by IDT in the Alt-R kit manual (1X RNP),4 and a 1:1 dilution with OptiMEM (0.5X RNP). This was done to evaluate RNP concentration-dependent effects on gene editing.

Cells were washed manually and using the Biomek automated method to ensure that fluorescence observed was due to tracrRNA incorporation into the cells. As expected, control cells that were not treated with ATTO 550 RNP did not fluoresce (Figure 5). Additionally, for both the manual and automated method, an RNP concentration-dependent increase in fluorescence was observed.

While the data for the 0.5X RNP condition was comparable between the manual and automated workflow, there was a demonstrable difference in the 1X RNP condition (Figure 5). The wells prepared using the Biomek i7 Hybrid were much more consistent, with a standard deviation of 3600 RFU. Wells that were treated by hand exhibited values ranging from approximately 5,000 to 30,000 RFUs and a standard deviation of 12,400 RFU. This highlights the utility of using automated liquid handlers for cell-based applications. The highest signal well observed in the manual samples was likely due to incomplete media removal during the cell washing step. The cell media at this step contained fluorescent tracrRNA, so incomplete media removal and cell washing could have led to an artificially high assay signal.

Additionally, the lowest signal observed in the manual wells was likely caused by inadvertent partial removal of the cell monolayer from the plate bottom. After reverse transfections, cell layers can be quite delicate, and aspirate/dispense steps must be designed to be very gentle. The Biomek can pipette at rates as slow as 1-2 μL/second and can be programmed to avoid pipetting in the center of the well. This precise and uniform pipetting is difficult to replicate by hand, likely leading to the highly variable results observed in this experiment. Taken together, this indicates that the fine tuning of liquid handling steps achievable using automated systems, like the Biomek i7 Hybrid workstation, can provide consistent results in adherent cell-based workflows.

T7 Endonuclease I (T7EI) Assay

The results presented above show that the automated method developed could reliably transfect HEK293 cells with RNP particles. In order to confirm that successful transfection lead to genome editing, the T7EI-based IDT Alt-R Genome Editing Detection Kit was used. In this protocol, the genomic region of interest was PCR-amplified from isolated gDNA. Following denaturation and slow renaturation of the PCR product, dsDNA heteroduplexes were formed because some PCR amplicons contained Hprt mutations and others did not, forming a DNA mismatch in the dsDNA. When the T7EI enzyme encountered this mismatch, it cleaved the DNA resulting in DNA fragments of various sizes. The IDT Alt-R CRISPR/Cas9 Control Kit was designed so that the Hprt region was amplified to create a fragment of approximately 1100 bp. If mutations were present, T7EI cleavage products of approximately 850 and 250 bp were expected.

For each condition tested, gDNA PCR amplicons from triplicate cell culture wells were combined into a single T7EI sample, thus the data presented in Figure 6 represents an average for each condition tested. Samples were prepared according to IDT protocols using the Biomek i7 Hybrid workstation, and DNA fragments were separated and quantified using a Bioanalyzer 2100 (Agilent) (Figure 6). As expected, independent of the method used, cells that were not treated with RNP gave rise to only a single band of the expected size, as only wild type Hprt was present (Lane 1,2). Similarly, cells treated with RNP containing a non-targeted crRNA produced only the same single band (Lane 10,11). Together these results demonstrated that the negative controls used here worked, as no editing was observed in either of these samples for either method.

Positive control wells contained either the IDT recommended amount of Hprt-targeted RNP (1X) or a 1:1 dilution of this condition (0.5X) to assess concentration-dependent effects on gene editing. As shown in Figure 6 and summarized in Table 1, gene editing was observed using both methods at both RNP conditions tested. A band corresponding to unedited Hprt homoduplexes was observed at 1100 bp, and cleavage products from heteroduplexes were clearly visible at 850 and 250 bp. These smaller bands were directly attributable to CRISPR/Cas9 mediated mutations and T7EI cleavage. In addition to this qualitative analysis, the Bioanalyzer allows semi-quantitation of gene editing (Table 1).

For every condition tested, ≥ 50% of the DNA was mutated. Further, the Biomek i7 Hybrid samples gave slightly higher editing efficiency compared to the manual experiments. One important caveat for quantitation of T7EI assay results, is that this assay will underestimate editing efficiency. During the heteroduplex formation step, two identical mutated DNA strands can form dsDNA. In this situation, the T7EI enzyme will not recognize a mismatch, and the DNA will not be cleaved. Additionally, 1 base pair insertions/deletions cannot be reliably detected by the T7EI assay, and some gRNAs will result in predominantly 1 base pair mutations. Thus, it remains likely that the values presented in Table 1 represent the lower limit of the amount of gene editing occurring in each well. Nevertheless, these data showed that the introduction of targeted RNPs using the Biomek i7 Hybrid workstation led to robust gene editing in HEK293 cells.

Summary

CRISPR/Cas9 gives researchers the ability to precisely edit the genomes of host species, which has allowed new insights to genotype-phenotype relationships. CRISPR has even become a primary method to interrupt gene function during genetic screening campaigns, replacing older RNAi-based techniques. While many methods exist to introduce functional RNP complexes into cells, lipofection represents the method most accessible to many labs. During lipofection, recombinant Cas9 protein is combined with a gRNA in a cationic lipid nanoparticle to generate functional RNPs, which are then introduced into living cells. This method is relatively fast and inexpensive and relies on commonly used tissue culture reagents, unlike nucleofection, which requires expensive pieces of specialty equipment and reagents.

The most straightforward way to perform CRISPR screening uses an arrayed strategy where each well of a microtiter plate contains gRNA(s) targeting a unique gene. This allows maximum flexibility, such as focused, pathway-specific screening campaigns, while providing data that is easily deconvoluted. Here we automated the delivery of RNP into HEK293 cells using IDT Alt-R reagents. The newly developed Biomek i7 Hybrid workstation produced data that compared favorably with assays performed by hand. Experiments using fluorescently labeled gRNA showed that transfection efficiency was comparable between the automated and manual workflow, but that the gentle pipetting techniques capable by the Biomek i7 Hybrid workstation method provided more reproducible, precise results.

Further, the automated method performed comparably to manual experiments regarding gene editing, as efficiencies of up to 60% were observed following IDT T7EI mismatch cleavage assays. The automated method also required less user hands-on time and reduced the chance of user-introduced pipetting errors. Taken together automated liquid handling using systems like the Biomek i7 Hybrid Automated Workstation provide great utility in workflows requiring high-throughput CRISPR/ Cas9 RNP delivery.

References

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Materials

Biomek Automated Workstations are not intended or validated for use in the diagnosis of disease or other conditions. This protocol is for demonstration only and is not validated by Beckman Coulter. Beckman Coulter makes no warranties of any kind whatsoever express or implied, with respect to this protocol, including but not limited to warranties of fitness for a particular purpose or merchantability or that the protocol is non-infringing. All warranties are expressly disclaimed. Your use of the method is solely at your own risk, without recourse to Beckman Coulter.