Abstract
The broad spectrum of activities displayed by CRISPR-Cas systems has led to biotechnological innovations that are poised to transform human therapeutics. Therefore, the comprehensive characterization of distinct Cas proteins is highly desirable. Here we expand the repertoire of nucleases for mammalian genome editing using the archetypal Streptococcus thermophilus CRISPR1-Cas9 (St1Cas9). We define functional protospacer adjacent motif (PAM) sequences and variables required for robust and efficient editing in vitro. Expression of holo-St1Cas9 from a single adeno-associated viral (rAAV) vector in the neonatal liver rescued lethality and metabolic defects in a mouse model of hereditary tyrosinemia type I demonstrating effective cleavage activity in vivo. Furthermore, we identified potent anti-CRISPR proteins to regulate the activity of both St1Cas9 and the related type II-A Staphylococcus aureus Cas9 (SaCas9). This work expands the targeting range and versatility of CRISPR-associated enzymes and should encourage studies to determine its structure, genome-wide specificity profile and sgRNA design rules.
INTRODUCTION
Clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated (Cas) proteins form a prokaryotic adaptive immune system and some of its components have been harnessed for robust genome editing1. Type II-based editing tools rely on a large multidomain endonuclease, Cas9, guided to its DNA target by an engineered single-guide RNA (sgRNA) chimera2 (See3, 4 for a classification of CRISPR-Cas systems). The Cas9-sgRNA binary complex finds its target through recognition of a short sequence called the protospacer adjacent motif (PAM) and subsequent base pairing of the guide RNA with the DNA to generate a specific double-strand break (DSB)1, 5. While Streptococcus pyogenes (SpCas9) remains the most widely used Cas9 variant for genome engineering, the diversity of naturally occurring RNA-guided nucleases is astonishing4. Hence, Cas9 enzymes from different microbial species can contribute to the expansion of the CRISPR toolset by increasing targeting density, improving activity and specificity as well as easing delivery1,6.
In principle, engineering complementary CRISPRCas systems from distinct bacterial species should be relatively straightforward, as they have been minimized to only two components. However, many such enzymes were found inactive in human cells despite being accurately reprogrammed for DNA binding and cleavage in vitro7–10. Nevertheless, the full potential of selected enzymes can be unleashed using machine learning to establish sgRNA design rules11,12.
Perhaps the most striking example of the value of alternative Cas9 enzymes is the implementation of the type II-A Cas9 from Staphylococcus aureus (SaCas9) for in vivo editing using recombinant adeno-associated virus (rAAV) vectors7, 13, 14. More recently, Campylobacter jejuni and Neisseria meningitidis Cas9s from the type II-C15 CRISPRCas systems have been added to this repertoire16, 17.
In vivo genome editing offers the possibility to generate phenotypes in animal models in order to better recapitulate the interactions between cell types and organs. In addition, it can be envisioned as a novel class of human therapeutics that enables precise molecular correction of genetic defects underlying diseases. As such, we have previously shown that rAAV- and zinc-finger nuclease (ZFN)-mediated liver targeting can correct disease phenotypes in neonatal and adult mouse models, a process currently under clinical investigation18–21. Therefore, further development of robust and wide-ranging CRISPR-based technologies for in vivo editing may help to decipher disease mechanisms and offer novel therapeutic options22, 23.
Here we revisited the properties of Streptococcus thermophilus type II-A CRISPR1-Cas9 system, a model nuclease of paramount importance to the entire CRISPR field, and engineered a potent RNA-guided nuclease for both in vitro and in vivo applications. S. thermophilus encodes up to two active type II-A systems (CRISPR1 and CRISPR3) that epitomize the instrumental role played by researchers focusing on the biology of CRISPR-Cas systems in the development of genome-engineering tools5, 24. In particular, study of the CRISPR1-Cas system has led to the following seminal discoveries; (i) spacer sequences are derived from phage genomes and plasmids25, (ii) CRISPR-Cas constitutes a bacterial adaptive immune system26, (iii) Cas9 is an endonuclease that cleaves DNA precisely 3 nucleotides upstream of the protospacer adjacent motif (PAM)27–29, (iv) sgRNA structural motifs govern function and orthogonality30, and (v) anti-CRISPR proteins (Acrs) block Cas9 activity31. The distinctive functional PAM sequences (NNAGAA and NNGGAA) of St1Cas9 increase the targeting flexibility and combinatorial potential of CRISPR-based genome editing tools. As shown for SaCas9, St1Cas9 can be efficiently packaged into an all-in-one rAAV for in vivo delivery to the liver. Moreover, we identified a family of anti-CRISPR proteins acting as off-switches for these two Cas9s. Currently, it is the only Cas9 functional in genome editing applications to be isolated from a nonpathogenic bacterium32.
RESULTS
Identification of an sgRNA architecture directing robust DNA cleavage by St1Cas9 in human cells
While characterizing the interplay between St1Cas9 and diverse Acr families isolated from virulent phages infecting S. thermophilus31, we were surprised by the substantial levels of editing achieved in human cells33. This observation contrasts with early reports indicating that this ortholog was mildly active7, 34. This led us to investigate whether we could further boost its activity. First, we added an N-terminal nuclear localization signal (NLS) to a previously described human codon-optimized expression construct35 and established a K562 cell line stably expressing St1Cas9 (S. thermophilus strain LMD-9) from the AAVS1 safe harbor locus36, 37 (Fig. 1a and Supplementary Fig. 1). St1Cas9 (1,121 aa) shares 17% and 37% identity with SpCas9 (1,368 aa) and SaCas9 (1,053 aa), respectively. Second, we adapted an sgRNA sequence used to monitor St1Cas9 activity in the heterologous host Escherichia coli30. We substituted a wobble base pair present in the lower stem of the repeat:anti-repeat region for a canonical Watson-Crick base pair in order to interrupt the RNA polymerase III termination signal (Fig. 1b). Then, we compared this sgRNA architecture (v1) to its counterpart containing a wild-type full length crRNA:tracrRNA duplex connected via a tetraloop (v0) by targeting FANCF, EMX1, and RUNX1 genes35 (Fig. 1c and Supplementary Fig. 1). St1Cas9-expressing human cells were transfected with increasing amounts of each sgRNA construct and the Surveyor nuclease assay was used to determine the frequency of indels characteristic of imprecise DSB repair by NHEJ36, 38
(Fig. 1d). The spectrum and frequency of targeted mutations was also analyzed using the complementary TIDE (Tracking of Indels by DEcomposition) method39 (Fig. 1d and Supplementary data set 1). Irrespective of the quantification method, the potency of sgRNA v1 was markedly superior. The increased activity was also observed when co-expressing St1Cas9 and its sgRNA transiently, a setting more typical of a genome editing experiment (Supplementary Fig. 1). This analysis revealed that high gene disruption rates could be obtained under standard conditions using St1Cas9 and a designed sgRNA in human cells.
Robust editing of target genes involved in liver metabolism by St1Cas9 in mouse cells
To further support these observations, we used CRISPOR40 to design several sgRNAs against Pck1, Pcsk9, and Hpd, three genes affecting liver function when disrupted. We reasoned that these targets would enable us to test the cleavage efficacy of St1Cas9 in vivo. When possible, we selected guides targeting essential protein domains and predicted to have few potential off-targets. Transient transfection of single vector constructs expressing both St1Cas9 and its sgRNA in mouse Neuro-2a cells revealed strong cleavage activity (18% to > 50% indels) at 14 out of 15 target sites highlighting the robustness of the system (Fig. 2a-c). Of note, this screen identified highly active sgRNAs targeting in the vicinity of mutations found in human HPD41, 42. Deficiency of 4-hydroxyphenyl-pyruvate dioxygenase (HPD), the second enzyme in the tyrosine catabolic pathway, causes Tyrosinemia type III (Orphanet ORPHA:69723) (Fig. 3a). Only three missense mutations are known to cause this rare disease (Prevalence <1/1,000,000) and we could target two of them with high efficacy (OMIM 276710) (Fig. 3c and Supplementary Fig. 2). Targeting the third mutation was not attempted due to the low specificity score of the guide. Taken together, these data suggest that St1Cas9 might enable in vivo genome editing if it could be packaged into a single rAAV particle alongside its sgRNA and the regulatory elements needed to drive its expression.
Potent in vivo genome editing using an all-in-one rAAV vector in newborn mice
Recombinant AAV vectors are prime in vivo gene delivery vectors for non-proliferative tissues as demonstrated in clinical trials for hemophilia B and inherited retinal dystrophy43–45. However, a limitation in the therapeutic use of rAAV is the loss of episomal vector genomes from actively dividing cells resulting in transient expression of therapeutic transgenes46–48. Hence, the combination of genome editing technology with rAAV-mediated delivery could lead to permanent genome modification and positive therapeutic outcome in young patients when tissues, such as the liver and retina, are still growing19, 49. As a side benefit, the elimination of vector genomes leads to transient nuclease expression in proliferating tissues that likely prevents accumulation of mutations at off-target sites19, 49.
To deliver holo-St1Cas9 (St1Cas9 + sgRNA) to the liver, we generated a hepatotropic rAAV serotype 813, 18–20 vector targeting Hpd exon 13 (aka AAV8-St1Cas9 Hpd G5) (Fig. 3c). To test the cleavage activity of St1Cas9 in vivo, we injected mice at day 2 of life into the retro-orbital sinus with increasing amounts of vector and isolated total liver DNA at day 28 post injection (Fig. 3b). The titration showed that the degree of target editing was substantial and dependent on the dose of AAV8-St1Cas9 (Fig. 3d).
To test if AAV8-St1Cas9 can lead to phenotypic correction in vivo, we used a mouse model of hereditary tyrosinemia type I (HT-I) (OMIM 276700) (Orphanet ORPHA:882), an autosomal recessive disease caused by a deficiency of fumarylacetoacetate hydrolase (FAH), the last enzyme of the tyrosine catabolic pathway (Fig. 3a). Of particular relevance to us, the incidence of HT-I reaches 1/1846 in a region of the province of Québec (Canada) while it is around 1/100,000 births worldwide50. Fah−/− mutant mice die as neonates with severe hepatic dysfunction and kidney damage due to the accumulation of toxic metabolites unless treated with nitisone (NTBC), a drug that inhibits Hpd upstream in the pathway (Fig. 3a)51.
Likewise, genetic ablation of Hpd in mice prevents liver damages and lethality leading to a much milder HT-III phenotype52, 53. Fah−/− mutant pups maintained on NTBC were injected at day 2 of life with AAV8-St1Cas9 Hpd G5 and then the drug was withdrawn shortly after weaning (Fig. 3b). Systemic delivery via a single neonatal injection rescued lethality in all mice while saline-treated animals had to be killed after ∼3 weeks as they met the weight loss criteria (Fig. 3e,f). Likewise, glycemia was normalized in the treatment groups (Fig. 3g). At low vector doses, we observed delayed but complete recovery, which is likely due to the potent selective growth advantage of targeted hepatocytes that can extensively repopulate the diseased organ51 (Fig. 3g). Notably, the excretion of succinylacetone, a toxic metabolite and a diagnostic marker for HT-I, was inversely correlated with the dose of rAAV demonstrating metabolic correction (Fig. 3h). These observations were recapitulated when targeting Hpd exon 8 at a site corresponding to a mutation also found in human patients (Supplementary Fig. 2). Therefore, rAAV-mediated delivery of St1Cas9 in vivo can correct a phenotype in neonatal mice by rewiring a metabolic pathway.
Lastly, we evaluated two additional vector architectures in order to minimize the size of rAAV and test the impact of the promoter on overall activity (Fig. 4). An rAAV vector (v3) containing an engineered liver-specific promoter (LP1b) combining the human apolipoprotein E/C-I gene locus control region (ApoE-HCR) and a modified human α1 antitrypsin promoter (hAAT) coupled to an SV40 intron and a synthetic polyadenylation element improved (>2 fold) cleavage efficacy at low rAAV dose as compared to the TBG promoter (Fig. 4a,b)54, 55. These modifications also led to the creation of a vector of ∼4.7 kb in size which is optimal for viral particle packaging13. Collectively, these data support the notion that St1Cas9 is a powerful tool for in vivo genome editing.
An anti-CRISPR family of proteins inhibits both SaCas9 and St1Cas9
Various families of anti-CRISPR proteins (Acrs) have been recently found in viruses and they block the activities of CRISPR-Cas systems in bacteria and archaea. Of interest here, these proteins could potentially be harnessed to fine-tune the activity of CRISPR-Cas9-based genome editing tools56. In theory, Acrs could be used as safety off-switches to block Cas9 activity in vivo and prevent overt off-target activity57. To begin to explore this avenue, we screened type II-A CRISPR-Cas9 inhibitor proteins (AcrIIAs) against SaCas9 and St1Cas931, 33, 58. Interestingly, the AcrIIA5 – family could inhibit both Cas9s in cultured cells (Fig. 5). AcrIIA4, an SpCas9 inhibitor57, 58, did not act on SaCas9 and St1Cas9 (Fig. 5). The broad range of activities displayed by these natural variants indicates that they could be engineered as highly potent Cas9 inhibitor for in vivo applications.
DISCUSSION
Here we report that St1Cas9 can be harnessed for robust and efficient genome editing in vitro and in vivo, thereby expanding the CRISPR-Cas toolbox. We optimized this previously overlooked system and validated its use in mice by demonstrating efficient rewiring and rescue of metabolic defects using all-in-one rAAV vectors. There is considerable interest in harnessing the diversity of Cas enzymes, but their implementation as genome editing tools is not a straightforward process7–10. Some enzymes simply fail to work and some choose their substrates promiscuously, necessitating thorough biochemical characterization59–65.Nevertheless, orthologous Cas9 enzymes with different PAM requirements increase targeting flexibility and allow multiplexing for combinatorial genetic screens, as demonstrated for SpCas9 and SaCas96, 11, 66. In that regard, St1Cas9 and SpCas9 also function orthogonally67. Moreover, sgRNAs for St1Cas9 and SaCas9 are not functionally interchangeable, which is likely due to their unique PAM specificity (Supplementary Fig. 3). These orthologous Cas9 systems could be adapted and combined for targeted knockout, transcriptional control, base editing, as well as “epigenome” editing1.
Further improvements in rAAV vector design; such as the use of alternative promoters, introns, and codon optimization schemes may yield to even higher activity in vivo13. The current size of the rAAV vector (v3) (Fig. 4b) is ∼4.7kb and, if needed, there is sufficient room to incorporate target sequences for the hematopoietic-specific microRNA miR-142 into the St1Cas9 expression cassette to prevent its expression in antigen-presenting cells (APCs) and induce robust tolerance to Cas913, 68–72.
Cas9 orthologs used for rAAV-mediated in vivo genome editing require a more complex PAM than the relatively simple NGG of SpCas91. This restricts the range of accessible targets but may reduce the occurrence of off-target mutagenesis73. The consensus PAM for St1Cas9 (LMD-9 and DGCC7710 S. thermophilus strains that differ by only 2 aa) has been defined as N1N2A3G4A5A6(W7), however sequences closely related to the consensus can be functional in test tubes and in bacterial cells27, 35, 74–77. While recognition of an A-rich PAM may ease targeting A/T-rich regions of genomes, we found that St1Cas9 can be targeted to both NNAGAA and NNGGAA PAMs in mammalian cells (Supplementary Fig. 4). Of note, in our limited dataset, the presence of an A at position 7 of an extended PAM correlates with high activity but is neither necessary nor sufficient (Supplementary Fig. 4 and Supplementary Tables 1–2). The length of the nonconserved linker (N1N2) has also been shown to be flexible and an extension from 2 to 3 bases is tolerated30, 78. However, we failed to reproduce this observation in human cells suggesting a higher stringency of the system in this context (Supplementary Fig. 4 and Supplementary Table 3). Structure-guided rational design and selection-based approaches could also enable St1Cas9 to recognize a broad range of PAM sequences35, 79, 80. Interestingly, making use of the natural diversity within S. thermophilus strains may achieve similar outcomes. For example, inferred consensus PAM sequences for St1Cas9 from strains CNRZ1066 and LMG13811 are NNACAA(W) and NNGCAA(A), respectively25, 32. Notably, the CRISPR1-Cas system of S. thermophilus strain LMG13811 transplanted in E. coli or reconstituted from purified components can target DNA using the NNGCAAA PAM78. At the protein level, the sequence of those three St1Cas9 variants diverges mostly within the C-terminal PAM-interacting (PI) domain, implying that they have evolved to recognize slightly distinct PAM sequences (Supplementary Figs. 5–6). Database mining predicts that even more diversity exists within CRISPR1-StCas9 systems81. Since the crRNA and tracrRNA are completely conserved30, we envisioned that a suite of tools based on naturally occurring St1Cas9 variants could be used in order to expand the targeting range of this genome editing system.
AUTHOR CONTRIBUTIONS
Conceptualization, S.C., D.A., A.D., and Y.D.; Methodology, S.C., D.A., A.D., J.L., and Y.D.; Investigation, S.C., D.A., A.D., J.L.; Writing – Original Draft, Y.D.; Writing – Review and Editing, S.C., D.A., A.D., and Y.D.; Supervision, Y.D.; Funding Acquisition, Y.D.
METHODS
Cell culture and transfection
K562 were obtained from the ATCC (CCL-243) and maintained at 37 °C under 5% CO2 in RPMI medium supplemented with 10% FBS, penicillin-streptomycin and GlutaMAX. Neuro-2a were obtained from the ATCC and maintained at 37 °C under 5% CO2 in DMEM medium supplemented with 10% FBS, penicillin-streptomycin and GlutaMAX. All cell lines are tested for absence of mycoplasma contamination. Cells (2E5 per transfection) were transfected using the Amaxa 4D-Nucleofector (Lonza) per manufacturer’s recommendations. K562 cell lines expressing SaCas9 and St1Cas9 from the AAVS1 safe harbor locus were generated as described36, 37. Briefly, simultaneous selection and cloning was performed for 10 days in methylcellulose-based semi-solid RPMI medium supplemented with 0.5 µg/ml puromycin starting 3 days post-transfection. Clones were picked and expanded in 96 wells for 3 days and transferred to 12-well plates for another 3 days before cells were harvested for western blot.
Genome editing vectors
Vectors for in vitro and in vivo genome editing with the CRISPR1-Cas9 (St1Cas9) system of S. thermophilus LMD-9 generated in this study are available from Addgene (Supplementary Fig. 7). The CRISPOR40 web tool was used to design guide (spacer) sequences against mouse and human targets (Supplementary Tables 1–4). A fraction of the guides targeting EMX1, FANCF, and VEGFA have been described previously35 (Supplementary Tables 1,4). DNA sequence for the spacers were modified at position 1 to encode a “G” due to the transcription initiation requirement of the human U6 promoter when required. Alternatively, the spacer length was increased to capture a naturally occurring “G”. The mammalian expression vector for St1Cas9 (LMD-9) fused to SV40 NLS sequences at the N- and C-terminus (MSP1594_2x_NLS; Addgene plasmid #110625) was constructed from MSP159435 (Addgene plasmid #65775, a gift from Keith Joung). The U6-driven sgRNA expression plasmids for St1Cas9 (LMD-9) (v1) (St1Cas9_LMD-9_sgRNA_pUC19; Addgene plasmid #110627) and SaCas97 (Supplementary Table 5) were synthesized as a gBlock gene fragments (Integrated DNA Technologies) and cloned into pUC19. BPK230135 (v0) (Addgene plasmid # 65778, a gift from Keith Joung) was used to compare St1Cas9 sgRNA architectures. The single vector mammalian expression system containing a CAG promoter-driven St1Cas9 LMD-9 and its U6-driven sgRNA (U6_sgRNA_CAG_hSt1Cas9_LMD9; Addgene plasmid #110626) was built from the above-described plasmids. The single vector rAAV-St1Cas9 LMD-9 systems containing liver-specific promoters (Supplementary Table 6) were assembled from the above-described components into a derivative of pX6027 (Addgene plasmid #61593, a gift from Feng Zhang) containing a deletion within the backbone to eliminate BsmBI restriction sites. The LP1b promoter was engineered by combining elements from previously described AAV expression cassettes54, 55. We deposited the most active version of this vector (v3) (pAAV_LP1B_St1Cas9_LMD-9_SpA_U6_sgRNA; Addgene plasmid # 110624). To establish clonal K562 cell lines constitutively expressing C-terminally tagged SaCas9 and St1Cas9 under the control of an hPGK1 promoter, the Cas9 ORFs from pX602 and MSP1594_2x_NLS were subcloned into AAVS1_Puro_PGK1_3xFLAG_Twin_Strep37 (Addgene plasmid # 68375). Untagged AcrIIA proteins were expressed transiently from a modified pVAX1 vector (Thermo Fisher Scientific) containing a beta-globin intron. The AcrIIA ORFs were codon-optimized for expression in human cells and synthesized as gBlock gene fragments (Integrated DNA Technologies) (Supplementary Table 7).
Surveyor nuclease and TIDE assays
Genomic DNA from 2.5E5 cells was extracted with 250 µl of QuickExtract DNA extraction solution (Epicentre) per manufacturer’s recommendations. The various loci were amplified by PCR using the primers described in Supplementary Table 8. Assays were performed with the Surveyor mutation detection kit (Transgenomics) as described36, 38. Samples were separated on 10% PAGE gels in TBE buffer. Gels were imaged using a ChemiDoc MP (Bio-Rad) system and quantifications were performed using the Image lab software (Bio-Rad). TIDE analysis was performed using a significance cut-off value for decomposition of p<0.00139
Recombinant adeno-associated virus production
Production of recombinant adeno-associated viral vectors was performed by the triple plasmid transfection method essentially as described82. Briefly, HEK293T17 cells were transfected using polyethylenimine (PEI, Polysciences) with helper plasmid pxx-680 (A gift from R.J. Samulski), the rep/cap hybrid plasmid pAAV2/8 (A gift from James Wilson) and the rAAV vector plasmid. Twenty-four hours post-transfection, media was replaced with growth media without FBS, and cells were harvested 24 hours later. rAAV particles were extracted from cell extracts by freeze/thaw cycles and purified on a discontinuous iodixanol gradient. Virus were resuspended in PBS 320 mM NaCl + 5% D-sorbitol + 0.001% pluronic acid (F-68), aliquoted and stored at − 80°C. rAAV were titrated by qPCR (Roche) using SYBR green and ITR primers as described83. The yields for all vectors varied between 1E13 and 2E13 vg/ml. The purity of the viral preparations was determined by SDS-PAGE analysis on a 10% stain free gel (Biorad) in Tris-Glycine-SDS buffer (Supplementary Figure 8). ITR integrity was assessed following a BssHII digestion of the AAV plasmid. The vector core facility at the Canadian neurophotonics platform (molecular tools) produced the rAAV8s.
Animal experiments
Fah−/− mice84 on a C57BL/6 genetic background were group-housed and fed a standard chow diet (Harlan #2018SX) with free access to food and water. Fah−/− mice drinking water was supplemented with 7.5 mg (2-(2-nitro-4-trifluoromethylbenzoyl)-1,3-cyclohexanedione) (NTBC)/L and pH was adjusted to 7.0. Mice were exposed to a 12:12-h dark-light cycle and kept under an ambient temperature of 23 ± 1 °C. Animals were cared for and handled according to the Canadian Guide for the Care and Use of Laboratory Animals. The Université Laval Animal Care and Use Committee approved the procedures.
Two days old neonatal mice were injected intravenously in the retro-orbital sinus85 with different doses of rAAV8 or saline in a total volume of 20 µL. Mice were weaned at 21 days of age and NTBC was removed 7 days later. Body weight and glycemia were monitored daily following NTBC removal. Mice were not fasted for measurement of glycemia, data collection occurred between 9–10 am. Animals were killed by cardiac puncture under anesthesia at predetermined time points or when weight loss reached 20% of body weight. Livers were snap frozen for downstream applications.
Urine collection and succinylacetone quantification
Urine from groups of 3–4 mice was collected overnight in metabolic cages (Tecniplast) 15 days after NTBC removal. Urine was centrifuged at 2000 rpm for 5 minutes, aliquoted and frozen at-80°C. Succinylacetone was quantified in urine samples by a sensitive method using gas chromatography–mass spectrometry (GC-MS) as previously described86. The biochemical genetics laboratory at the centre hospitalier universitaire de Sherbrooke performed the analyses.
ACKNOWLEDGMENTS
This study was supported by grants from the Canadian Institutes of Health Research (CIHR) and the Banting Research Foundation to Y.D. Salary support was provided by the Fonds de la recherche du Québec-Santé (FRQS) to Y.D. D.A. holds a Vanier Canada graduate scholarship. Partial salary support to A.D. was provided by a Desjardins scholarship from the Fondation du CHU de Québec. We thank Marie-Ève Paquet and the skilled vector core facility staff at the Canadian neurophotonics platform (molecular tools) for rAAV8 production, as well as Paula Waters and Denis Cyr for their high level analytical expertise on the quantitation of metabolites. Robert Tanguay provided the mouse model of HT-I, nitisone, expertise and support. Sylvain Moineau ignited our interest in S. thermophilus CRISPR systems and graciously shared his insights on CRISPR-Cas biology and Acrs.
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