Genetic knockdown of genes that are obscure, conserved and essential using CRISPR interference methods in the fission yeast S. pombe

CRISPR-Cas (for ‘clustered regularly interspaced short palindromic repeat–CRISPR-associated’) is a form of adaptive immunity that restricts viral infection in bacteria and archaea (Knott and Doudna, 2018). In laboratories, artificial type II CRISPR-Cas system cleave specific target DNAs with a ribonucleoprotein complex comprising the Cas9 DNA endonuclease and small guide RNA (sgRNA). sgRNA has a targeting sequence (formally called a spacer sequence), which is a 20–30-nt sequence complementary to a specific target DNA (Jinek et al., 2012). The Cas9 ribonucleoprotein complex recognizes a specific target DNA through two mechanisms, Watson–Crick base-pairing with the targeting/spacer sequence by the sgRNA, and 2–5-bp PAM (protospacer adjacent motif) sequence recognition by Cas9 protein (PAM is situated immediately to the 3′-side of the targeted, protospacer DNA sequence) (Sternberg et al., 2014).

Given that this artificial CRISPR-Cas system contains only cas9 and sgRNA genes, it is easily manipulated so as to cleave arbitrary target DNAs by altering the targeting sequence encoded by the sgRNA gene; therefore, Cas9 and sgRNA are utilized for genome editing in a broad range of organisms, including Schizosaccharomyces pombe (Hayashi and Tanaka, 2019; Jacobs et al., 2014; Rodríguez-López et al., 2016; Zhang et al., 2018). In addition to genome editing, this artificial system is utilized for transcriptional alteration. Catalytically inactive Cas9 (dCas9) retains specific DNA-binding activity, and its binding to specific DNA targets can inhibit transcription there (Bikard et al., 2013; Qi et al., 2013). This technology, CRISPR interference (CRISPRi), was originally established using bacterial systems, and later implemented in many other organisms. For fission yeast, Zhao and Boeke established Cas12a (class 2 type V system)-mediated CRISPRi (Zhao and Boeke, 2020), and we recently implemented the Cas9-mediated method (Ishikawa et al., 2021). Transcriptional repression efficiency of CRISPRi is affected by the position at which dCas9 binds to the target DNA, and the effective binding position differs for each organism (Bikard et al., 2013; Gilbert et al., 2014; Lawhorn et al., 2014; McGlincy et al., 2021; Qi et al., 2013). We revealed that in S. pombe, complementary binding by the targeting sequence to either non-template strand at the transcription initiation (start) site (TSS) of a target gene or template strand 60–120 bp downstream of the TSS is preferred for efficient transcriptional repression by dCas9-mediated CRISPRi (Ishikawa et al., 2021). This rule for selecting targeting sequences can dramatically reduce the labor needed to design effective sgRNAs for dCas9-mediated CRISPRi in S. pombe.

The fission yeast S. pombe is a well-studied model organism. Genetic analyses using it have revealed numerous conserved gene functions among eukaryotes. However, characterizing genes essential for cellular viability is still challenging, given that perturbing functions of essential genes causes cell death. Thus, construction of strains defective in essential genes is not feasible. Physiological functions of essential genes have traditionally been analyzed using conditionally defective mutants, such as temperature-sensitive mutants, but these are not always available. Although recently developed methods, such as forced protein degradation, promoter replacement and 3′ untranslated region (3′UTR) disruption, enable researchers to reduce expression and function of target genes without using conditionally defective mutations, these methods are time-consuming, as they require modification of genomic DNA sequences of target genes (Bähler et al., 1998; Kanke et al., 2011; Roguev et al., 2008; Schuldiner et al., 2005). CRISPRi is another feasible option to conditionally repress transcription of target genes. In contrast to other methods, dCas9-mediated CRISPRi can be executed simply by introducing a plasmid harboring a sgRNA gene and the dCas9 gene into S. pombe cells and inducing expression of dCas9 (Fig. 1A). In the plasmid, pSPdCas9, the dCas9 gene is placed under the regulatable promoter, nmt1-41p. Genes can be knocked down arbitrarily using this method at desired timing, as long as a proper sgRNA is designed (Ishikawa et al., 2021). Owing to such simplicity of implementation, dCas9-mediated CRISPRi is potentially suitable for systematic functional analyses of essential genes in S. pombe. However, to our knowledge, this technology had not been utilized for this purpose.

Here, we successfully improved dCas9-mediated CRISPRi in fission yeast to achieve more efficient transcriptional repression and broader options for sgRNA design. Using the original and improved versions of CRISPRi, we then systematically knocked down seven genes that are evolutionarily conserved from yeasts to mammals but have not been well characterized. This allowed us to investigate phenotypes that emerged after their perturbation in S. pombe. CRISPRi is thus a feasible option to study physiological roles of essential genes for which conditional mutant strains are not available.

Although conventional CRISPRi, which consists of dCas9 protein and an sgRNA against a target gene, can repress transcription of a model gene, ade6⁺, by as much as ∼85% in S. pombe (Ishikawa et al., 2021), simultaneously applying two distinct sgRNAs against a single target gene reportedly increased transcriptional repression efficiency of CRISPRi in E. coli (Qi et al., 2013). This is called double sgRNA repression. As each fission yeast gene has two distinct spots potentially suitable for sgRNAs in close proximity to the TSS and 60–120 bp downstream (Ishikawa et al., 2021), by targeting both spots, we examined whether double sgRNA repression can enhance transcriptional repression by CRISPRi in this organism. To implement double sgRNA repression for fission yeast, we constructed a plasmid that has two sgRNA units in addition to a dCas9 gene placed under the thiamine-repressible nmt1-41p promoter (Fig. 1B, pSPdCas9.5). We then inserted two effective targeting sequences, ‘a5’ and ‘a10’, into the sgRNA units on this plasmid. Sequence a5 binds in the vicinity of the TSS of the ade6⁺ gene, whereas a10 binds downstream of the TSS (Fig. 2A). The resulting plasmids, pSC5(a5,a10) and pSC5(a10,a5), were introduced to perform double sgRNA repression of ade6⁺ gene expression.

In cells harboring a plasmid containing either a5 or a10 sgRNA [pSC2(a5), pSC2(a10), pSC5(a5,ns) or pSC5(ns,a5)], levels of ade6⁺ mRNA decreased to ∼16% of that in control cells with a nonsense sgRNA (ns), consistent with results previously reported (Ishikawa et al., 2021) (Fig. 2B). In cells harboring a plasmid expressing both a5 and a10 sgRNAs simultaneously [pSC5(a5,a10) and pSC5(a10,a5)], ade6⁺ mRNA abundance was further reduced to 8–9% of the control, indicating that double sgRNA repression reduced the ade6⁺ mRNA level by half in comparison with conventional CRISPRi using a single sgRNA. In cells with reduced ade6⁺ gene expression, colonies became reddish due to accumulation of a red pigment, 5-aminoimidazole ribotide, which is an intermediate of adenine biosynthesis (Fig. 2C). Consistent with lower expression of ade6⁺, the colony color of cells subjected to double sgRNA repression was darker than of cells subjected to conventional CRISPRi with a single sgRNA. Greatly enhanced transcriptional repression efficiency by CRISPRi by double sgRNA repression in S. pombe was thus confirmed.

The PAM sequence, which is recognized by Cas9 protein, is a main limitation for designing sgRNAs. The dCas9 utilized in our CRISPRi method is derived from Streptococcus pyogenes. The PAM of S. pyogenes Cas9 is 5′-NGG (N=A, C, G or T) (Jinek et al., 2012). Therefore, targeting sequences for dCas9-mediated CRISPRi must be located right next to this sequence motif. Although a reverse targeting sequence binding at the TSS of a target gene and a forward targeting sequence binding ∼90 bp downstream of the TSS are most preferable for transcriptional repression by dCas9-mediated CRISPRi in S. pombe (Ishikawa et al., 2021), targeting sequences are not always available at such positions because of the limitation imposed by the PAM sequence. Recently, a variant of Cas9 that recognizes relaxed PAM sequences, 5′-NGN, was artificially created, and successfully applied to genome and base editing (Walton et al., 2020). To investigate whether this variant, designated as SpG, is also suitable for CRISPRi, we generated plasmids pdSpG.2 and pdSpG.5 in which nucleotide changes of the SpG were introduced to the dCas9 gene (Fig. 1C,D). Eight targeting sequences were designed at the illustrated positions in the ade6⁺ gene (Fig. 2D), inserted into the sgRNA cassette of pdSpG.2, and resulting plasmids were introduced into S. pombe cells for evaluation of their transcriptional repression capacity (Fig. 2E,F). Among targeting sequences tested (sa1–sa8), sa2, which binds to the non-template strand in the vicinity of the TSS, showed the most effective repression. In this strain, the ade6⁺ mRNA level was reduced to 12.1% of the control (Fig. 2E), and cells formed reddish colonies (Fig. 2F). These results indicate that dSpG-mediated CRISPRi is as effective as dCas9-mediated CRISPRi, and that it is particularly useful in cases in which targeting sequences linked to canonical PAM sequences for dCas9, 5′-NGG, are not found in regions preferred for transcriptional repression.

Notably, dSpG is not equally effective for targets with each sequence of the 5′-NGN (Walton et al., 2020). Consistent with a previous report (Walton et al., 2020), PAM sequences of 5′-DGHG (D=A, G or T; H=A, C or T) yielded the most effective transcriptional repression of the ade6⁺ gene (Fig. 2E,G). These results confirmed that the preferred PAM sequences (5′-DGHG) should be chosen for effective CRISPRi with the dSpG. Although dSpG provides additional variation of PAM sequences available for CRISPRi, targets with 5′-NGG PAM still have to be repressed by dCas9 rather than dSpG, given that some options of 5′-NGG are not preferred by SpG (Walton et al., 2020) (Fig. S1).

S. pombe is an excellent unicellular model organism to study molecular and physiological functions of evolutionarily conserved genes. Among 1221 essential genes, deletion of which causes cell death, ∼20 remain uncharacterized in this organism, whereas the others have been characterized experimentally and/or have had functions inferred based on sequence homology (Pombase, 12/09/2020, https://www.pombase.org/). Seven of these uncharacterized genes are evolutionarily conserved from yeasts to mammals, and here they are designated as conserved essential obscure (ceo) genes. According to their positions in the fission yeast genome, ceo genes were systematically numbered from ceo1⁺ to ceo7⁺ (Table 1). Three of the seven ceo genes (ceo2⁺, ceo5⁺ and ceo7⁺) are essential, even in human cell lines (Blomen et al., 2015; Wang et al., 2015), and another (ceo6⁺) has been predicted as an essential human gene (Georgi et al., 2013). Human homologs of the five ceo genes (ceo1⁺, ceo2⁺, ceo3⁺, ceo6⁺ and ceo7⁺) are thought to be involved in diseases when they malfunction (Belyaeva and Kedishvili, 2002; Blomen et al., 2015; Curbo et al., 2006; Fatima et al., 2021; Guimier et al., 2016; Matsuura et al., 2000; Pastore et al., 2021; Wang et al., 2009, 2015; Wilcox et al., 2007) (Table S1). Thus, revealing the functions of these genes might not only uncover unidentified physiological processes essential for cell viability, but might help to improve human health. Given that conditional mutants of these ceo genes have not been isolated to date, we studied their physiological roles using CRISPRi techniques.

First, ceo genes were knocked down using conventional CRISPRi with dCas9 and a single sgRNA. According to a previous report (Ishikawa et al., 2021), sgRNAs were designed to bind close to the TSS of the target ceo gene or ∼90 bp downstream of the TSS (Fig. 3A–C). Fission yeast cells harboring a pSPdCas9-derived plasmid with one of the designed sgRNAs were cultivated in the presence of 20 μM thiamine, which represses expression of dCas9 from the nmt1-41 promoter, and then placed in medium without thiamine to induce dCas9 expression. As all ceo genes are essential for cell viability, knockdown of their expression was expected to inhibit colony formation. Consistent with this prediction, colony formation was clearly inhibited in ceo3⁺ and ceo5⁺ knockdown strains with two and three designed sgRNAs, respectively (Fig. 3D,E). Colony formation of a ceo7⁺ gene knockdown strain was slightly inhibited (Fig. 3F). These observations demonstrate that dCas9-mediated CRISPRi is applicable for conditional knockdown of these ceo genes.

mRNA levels of ceo genes were examined in these CRISPRi knockdown strains. Cells were cultivated in the absence of thiamine for 26 h and mRNA levels were quantified by RT-qPCR (Fig. 3A–C). Consistent with inhibition of colony formation, the abundance of ceo3⁺ mRNA was reduced by half (50–54%) in ceo3⁺ knockdown cells harboring ceo3-1 or ceo3-2 sgRNA (Fig. 3A,D). Likewise, ceo5-1, ceo5-2 and ceo5-4 sgRNAs greatly reduced the level of ceo5⁺ mRNA to 29–36% of the control, and ceo7-7 sgRNA reduced ceo7⁺ mRNA abundance to 54% of the control (Fig. 3B,C,E,F). Intriguingly, mRNA abundance and cell viability did not appear to correlate linearly. For example, ceo3⁺ mRNA levels were reduced to a similar extent by ceo3-1, ceo3-2 and ceo3-5 sgRNAs, but the extent of inhibition of colony formation by these sgRNAs differed greatly (Fig. 3A,D). ceo3-1 and ceo3-2 sgRNAs blocked colony formation nearly completely, whereas ceo3-5 did not. Similarly, ceo5-1, ceo5-2 and ceo5-4 sgRNAs inhibited colony formation, but ceo5-3 did not, although these four sgRNAs repressed transcription of the ceo5⁺ gene to a similar extent (Fig. 3B,E). These observations suggest that each gene might have a threshold value for reduction of the mRNA level, below which cell proliferation abruptly diminishes.

As described above, conventional dCas9-mediated CRISPRi using one sgRNA was efficient enough to reduce colony formation when employed to knockdown ceo3⁺, ceo5⁺ and ceo7⁺ gene expression. However, this method did not inhibit colony formation when applied to ceo1⁺, ceo2⁺, ceo4⁺ and ceo6⁺ (Figs 4, 5; Fig. S2). These results suggest that efficiency of transcriptional repression by the conventional method was not sufficient to impair colony formation. To enhance transcriptional repression, we next utilized double sgRNA repression CRISPRi to knockdown expression of these genes, and expression of ceo1⁺ and ceo6⁺ was successfully repressed by combined use of two sgRNAs (Fig. 4). Four sgRNAs were designed for repression of ceo6⁺ expression. ceo6-1 binds to the non-template strand at the TSS, whereas ceo6-5, ceo6-6 and ceo6-3 bind to the template strand ∼90 bp from the TSS (Fig. 4A). Although both ceo6-1 and ceo6-3 sgRNA alone could reduce the ceo6⁺ mRNA level by ∼50%, neither could inhibit colony formation (Fig. 4A,C). When these two sgRNAs were applied in combination, the ceo6⁺ mRNA level decreased further and colony formation was inhibited (Fig. 4A,C). To repress ceo1⁺ gene expression, four sgRNAs, ceo1-9, ceo1-10, ceo1-11 and ceo1-12, were designed as illustrated in Fig. 4B. Although each of these sgRNAs individually could reduce the ceo1⁺ mRNA level to ∼14% of that in the control with a nonsense sgRNA, combination of ceo1-9 and ceo1-11 further reduced the level to 4.8%. Given that a single vegetative cell normally contains 0.81 molecules of ceo1⁺ mRNA (Marguerat et al., 2012), the result indicated that double repression with ceo1-9 and ceo1-11 sgRNAs reduced the ceo1⁺ mRNA abundance to ∼0.04 molecule per cell, although colony formation was apparently not inhibited (Fig. 4D). The threshold ceo1⁺ mRNA level for cell proliferation is thus extremely low.

dSpG-mediated CRISPRi was applied to knockdown ceo4⁺ expression, as sgRNAs for dCas9 could not be designed at exactly the preferred spots due to the limitation of the PAM sequence (Fig. 5A). Together with three sgRNAs for dCas9, ceo4-5, ceo4-6 and ceo4-7, three more sgRNAs for dSpG, ceo4-8, ceo4-9 and ceo4-10, which bind closer to the preferred spots than those for dCas9 do, were examined for transcriptional repression. Five of the seven sgRNAs tested here reduced the ceo4⁺ mRNA level by ∼70%. Double sgRNA repression by combining ceo4-9 and ceo4-10 sgRNAs further reduced the mRNA level to 21.5% of the control, which corresponds to ∼0.67 molecule per cell (Marguerat et al., 2012) (Fig. 5A), although it apparently did not affect colony formation (Fig. 5B). As with ceo1⁺, the threshold of the ceo4⁺ mRNA level for cell proliferation is low.

The ceo2⁺ gene has at least five TSSs in a 25-bp region of its 5′UTR (Meylan et al., 2020), which makes it difficult to design sgRNAs that bind at the TSS or to ∼90 bp downstream of the TSS. As attempts to repress ceo2⁺ transcription with dCas9- or SpG-mediated CRISPRi with a sgRNA that binds near these preferred spots (ceo2-3, ceo2-4 and ceo2-10) were unsuccessful, we sought for effective sgRNAs outside these spots (Fig. S2). Among those tested, three sgRNAs for dCas9 (ceo2-1, ceo2-12 and ceo2-13) reduced the ceo2⁺ mRNA level by 30–40% (Fig. S2B). Double sgRNA repression using a combination of ceo2-1 and ceo2-12 further reduced the mRNA level by 55%, although colony formation was not affected (Fig. S2B,C). Of note, the normal abundance of ceo2⁺ mRNA is estimated to be as low as 1.3 molecules in a single vegetative cell (∼0.58/cell after CRISPRi) (Marguerat et al., 2012).

In summary, colony formation of strains knocked down on the ceo1⁺, ceo2⁺ or ceo4⁺ gene was indistinguishable from that of the nonsense control, although their transcription levels were reduced to less than one molecule per cell. It is possible that their mRNA and protein molecules have such long half-lives that a sufficient amount of protein was synthesized and accumulated from a small number of mRNA molecules; thus, their abundances were hardly reduced by CRISPRi repression.

As described above, we established ceo gene knockdown strains using CRISPRi. mRNA levels of targeted ceo genes in these strains decreased by 48.6% to 94.3% (Table 1). ceo1⁺, ceo2⁺ and ceo6⁺ gene expression was reduced most efficiently by double sgRNA repression dCas9-CRISPRi with combinations of ceo1-9 and ceo1-11, ceo2-1 and ceo2-12, and ceo6-1 and ceo6-3 sgRNAs, respectively. Gene expression of ceo3⁺, ceo5⁺ and ceo7⁺ was repressed most efficiently by conventional dCas9-CRISPRi using ceo3-2, ceo5-4 and ceo7-7 sgRNAs, respectively. ceo4⁺ gene expression was knocked down by the dSpG-mediated double sgRNA repression with ceo4-9 and ceo4-10 sgRNAs. Thus, knockdown strains with conventional or modified CRISPRi were used in the following experiments. Four out of the seven knockdown strains (ceo3⁺, ceo5⁺, ceo6⁺ and ceo7⁺) showed inhibited colony formation (Table 1, Fig. 6A,B; Fig. S3A). To prove that dCas9-mediated CRISPRi is applicable for characterizing physiological roles of essential genes, maximum growth rate (MGR, divisions/hour) in liquid culture and cellular morphology were analyzed in cells in which ceo gene expression was repressed.

Inhibition of cell proliferation is reflected as a decrease of the MGR, which is the reciprocal of the minimum time required for cell density to double. The MGR was calculated by measuring the optical density at 600 nm (OD₆₀₀) of the cell culture as follows. After 50 h incubation in liquid Edinburgh minimal medium 2 (EMM2) without thiamine for induction of CRISPRi, ceo knockdown, cells were resuspended in fresh EMM2 and placed in a spectrometer equipped with a shaker and a heater. OD₆₀₀ of the cell culture was then measured every 30 min at 33°C (Fig. 6C,D, Fig. S3B). Growth rate (divisions/hour) was calculated every nine measurements (4 h). Among growth rates throughout an experiment, the maximum value was defined as the MGR (Fig. 6E, Fig. S3C; see Materials and Methods for details). MGRs of ceo-knockdown strains (ceo-KD strains) correlate with their colony formation ability on solid medium. For example, ceo3⁺ knockdown reduced MGR to 0.10 divisions/hour, whereas control cells grew at the MGR of 0.21 (Fig. 6E), consistent with severe growth retardation on solid medium by ceo3⁺ gene knockdown (Fig. 6A). Knockdown of ceo7⁺ gene expression, which resulted in only a slight decrease of colony formation on solid medium, apparently did not affect the MGR (Fig. 6A,E). It is of note that MGR might be overestimated because there can be an increase of OD caused by cellular elongation without cell division observed in knockdown strains of ceo5⁺ and ceo6⁺ as described below.

To examine how ceo gene knockdown affects cell cycle progression and cell shape, S. pombe cells were fixed with glutaraldehyde before and after CRISPRi knockdown and stained with DAPI, which fluorescently labels nuclear DNA and surfaces of fixed fission yeast cells (Fig. 7). Control cells harboring a vector plasmid with nonsense sgRNA gene(s) showed normal cellular morphology before and after induction of dCas9 from the thiamine-repressive promoter. This indicates that expression of dCas9 and nonsense sgRNA does not affect cell shape or mitotic nuclear division (Fig. 7; Fig. S4). All ceo gene knockdown cells showed normal cell and nuclear morphology in the presence of thiamine, which represses expression of dCas9. After removal of thiamine for induction of dCas9, the ceo5⁺ and ceo6⁺, knockdown cells showed an elongated cell shape. As these elongated cells largely contain single nuclei, knockdown of the ceo5⁺ and ceo6⁺ genes might result in blocked or delayed cell cycle progression before mitotic nuclear division (Nurse et al., 1976). On the other hand, ceo3⁺ knockdown cells were short and slightly round, and they frequently showed plasmolysis-like morphology, in which cell surface materials strongly stained with DAPI accumulated and intruded into the cytoplasm. The ceo3⁺ gene product might be required for maintenance of cell wall and/or cell membrane integrity. Collectively, CRISPRi enabled conditional knockdown of essential ceo genes and phenotypic analyses of knockdown cells. CRISPRi is a useful tool to characterize physiological roles of essential genes in S. pombe when their conditional lethal mutants are not available.

Analyzing the functions of essential genes in S. pombe has been challenging. This study showed that CRISPRi is a feasible option to resolve this issue by providing a systematic strategy to selectively knockdown expression of essential genes. Conventional dCas9-mediated CRISPRi successfully inhibited transcription of ceo genes efficiently enough to inhibit cell proliferation of knockdown strains of ceo3⁺, ceo5⁺ and ceo7⁺. The double sgRNA repression technique implemented in this study enhances transcriptional repression efficiency, and successfully repressed ceo6⁺ gene expression to inhibit cell proliferation. dSpG-mediated CRISPRi is also useful, particularly in cases in which canonical PAM sequences of dCas9 (5′-NGG) are not available at preferred target gene sites. ceo4⁺ gene expression was reduced by 75% using dSpG in combination with the double sgRNA repression technique. To prove that CRISPRi is applicable for functional characterization of essential genes, physiological roles of ceo genes were analyzed by utilizing the gene knockdown strains constructed above – knockdown of ceo5⁺ and ceo6⁺ genes resulted in cell elongation, whereas ceo3⁺ knockdown caused cell shortening and a plasmolysis-like phenotype, implying that these ceo genes might be involved in cell cycle control and/or maintenance of cell shape integrity. Thus, CRISPRi provides an alternative means of characterizing function of essential genes in S. pombe when suitable conditional lethal mutant alleles are not available.

Based on our experience with systematic knockdown of ceo genes by CRISPRi in this study, we propose a protocol to construct CRISPRi knockdown strains as follows. (1) Test whether conventional dCas9-mediated CRISPRi with a single sgRNA, which preferably binds at the TSS or ∼90 bp downstream of the TSS, can sufficiently repress expression of the target gene. (2) If expression of the target gene is not sufficiently repressed by conventional CRISPRi, the double sgRNA repression method should be considered. (3) If the mRNA level of the target gene is not reduced even with double sgRNA method, dSpG-mediated CRISPRi should be employed with either a single or double sgRNAs.

During construction of ceo gene knockdown strains, we obtained a series of strains in which mRNA levels of the ceo genes decreased to various extents, and we found that subtle differences in the mRNA level often resulted in large differences in cell proliferation. Although we do not know whether this abrupt reduction of cell proliferation at a particular level of transcription holds true for all essential genes, it would be expected that there is a threshold mRNA level for each essential gene that is required to support normal cell proliferation. Knockdown strains constructed in this study allowed us to calculate such a threshold, designated as the ‘sufficient transcription index’ for ceo genes (Table 1). The sufficient transcription index was calculated as the average of the minimal transcription level that supported normal colony formation and the maximum transcription level that failed to support normal colony formation. Among ceo genes, the highest value of the sufficient transcription index was 55.8% (ceo7⁺), and this suggests that essential genes are often functional even when their transcription rate decreased to ∼60% of normal. Therefore, when CRISPRi for an essential gene does not decrease the mRNA to less than 60%, it is worth seeking conditions that improve repression efficiency by using double sgRNA repression and/or dSpG, to develop a conditionally defective strain.

Among human homologs of the seven ceo genes, six genes are involved in cancer cell viability and/or human diseases (Table S1) (Belyaeva and Kedishvili, 2002; Blomen et al., 2015; Curbo et al., 2006; Fatima et al., 2021; Georgi et al., 2013; Guimier et al., 2016; Matsuura et al., 2000; Pastore et al., 2021; Wang et al., 2009, 2015; Wilcox et al., 2007), implying that their essential roles are conserved evolutionarily from fission yeast to humans. Although the precise molecular functions of these gene products remain largely unknown, morphological defects caused by CRISPRi gene knockdown strongly suggest that ceo5⁺ and ceo6⁺ function in normal cell cycle progression, and that ceo3⁺ helps to maintain the integrity of cellular structures (Fig. 7). Interestingly, a high-throughput study showed that overproduced fluorescently labeled Ceo6 protein was localized at the spindle pole body in S. pombe (Matsuyama et al., 2006); thus, Ceo6 protein might contribute to formation of the mitotic spindle, which is essential for chromosome segregation. The same report found that Ceo3 protein was localized on the cell surface and as cytoplasmic dots, and this is consistent with our hypothesis that Ceo3 is involved in maintaining integrity of cell surface structure (Matsuyama et al., 2006). Detailed studies of Ceo proteins will help reveal how their human orthologs contribute to human health, and the CRISPRi strains constructed here are useful resources for such studies.

The present study shows that CRISPRi is a feasible option to characterize essential gene functions in S. pombe. Plasmid-based CRISPRi for fission yeast is easy to handle for knockdown of essential genes. This method is suitable for high-throughput applications such as construction of a genome-wide S. pombe gene knockdown strain library, which complements the haploid gene deletion strain library covering the only non-essential genes (Kim et al., 2010). Thus, CRISPRi in fission yeast will expand accessibility to genetic analyses for functional characterization of essential genes and greatly advance understanding of their conserved functions among eukaryotes.

Growth media and yeast culture conditions were as previously described (Moreno et al., 1991). Edinburgh Minimal Medium 2 (EMM2) contains 2% glucose. EMM2 supplemented with 20 µM thiamine (EMM2+T20) was utilized to repress CRISPRi of fission yeast strains. A low-adenine medium to detect repressed ade6⁺ gene expression was prepared by supplementing EMM2 with 7.5 mg/l adenine. Fission yeast (S. pombe) strains carrying a plasmid for CRISPRi were prepared by transformation of strain sp685 (h⁻ leu1⁻) using the standard lithium acetate method.

The pSPdCas9.5 plasmid was constructed by modifying the pSPdCas9 plasmid, which contain a sgRNA gene and a dCas9 gene (Ishikawa et al., 2021). To insert an additional sgRNA gene into pSPdCas9, a cloning site was inserted into it by ligating a double-stranded oligo-DNA containing restriction sites (PmeI and BamHI) at the plasmid SbfI site. The resulting plasmid with this additional cloning site was named pSPdCas9.2 (Fig. 1A). A PaqCI-site version of the sgRNA gene (sgRNA2) was constructed by inserting a double-stranded oligo-DNA containing two PaqCI sites into the BbsI-cleaved sgRNA gene on pSPdCas9. This plasmid (pSPdCas9.3) was utilized as a template to amplify a DNA fragment containing the sgRNA2 by polymerase chain reaction (PCR) with primers KI1169 and KI1248 (see Table S2 for primer sequences). The PCR product was cleaved with BamHI and was then ligated into the PmeI-BamHI site of the pSPdCas9.2. The resulting plasmid containing two sgRNA genes was named pSPdCas9.5 (Fig. 1B).

The SpG version of pSPdCas9.2, which was named pdSpG.2 (Fig. 1C), was constructed as follows. A 1553-bp DNA fragment, including SpG mutations that cause amino acid substitutions D1135L, S1136W, G1218K, E1219Q, R1335Q and T1337R, was amplified from plasmid pCAG-CBE4max-SpG-P2A-EGFP (Addgene 139998) by PCR with primers KI1423 and KI1424 (Walton et al., 2020). This DNA fragment was integrated into the corresponding region of pSPdCas9.2 by recombination with another DNA fragment amplified from pSPdCas9.2 by PCR with KI1425 and KI1426 (Nozaki and Niki, 2019). Given that the latter fragment contains mutations that cause amino acid substitutions at catalytic residues (D10A and H840A), the SpG encoded in pdSpG.2 is catalytically inactive (we call this mutant protein dead SpG or dSpG). pdSpG.2 was further modified to construct another plasmid for dSpG-mediated, double sgRNA repression (pdSpG.5, Fig. 1D), by inserting the DNA fragment containing the sgRNA2 to the pdSpG.2 in the same way as for pSPdCas9.5 construction.

Modifications of targeting sequences of sgRNAs on plasmids used in this study were conducted as previously reported (Ishikawa et al., 2021). Genomic positions of TSSs utilized to design targeting sequences of sgRNAs were previously determined (Li et al., 2015; Meylan et al., 2020). 5′ UTR regions indicated in this study were obtained from Pombase (https://www.pombase.org/). DNA sequences of targeting sequences and primers are available in Table S2. A plasmids list with brief descriptions on their structures is available in Table S3.

dCas9-mediated CRISPRi used in this study is conditionally inducible (Ishikawa et al., 2021). Transcription of the dCas9 gene is controlled by the nmt1-41p promoter, which is fully inhibited in the presence of 15 µM thiamine in media (Forsburg, 1993). Removing thiamine from the media induces dCas9 transcription and then CRISPRi. This procedure was conducted as previously reported (Ishikawa et al., 2021). Briefly, before induction of CRISPRi, fission yeast strains were grown on EMM2 plates supplemented with 20 µM thiamine at 33°C for about 24 h. To induce CRISPRi, cells were extensively washed and were subjected to culture in EMM2 liquid medium without thiamine for 6 h. This culture was diluted to 9.4×10⁴ cells/ml in EMM2 medium without thiamine, followed by incubation at 33°C for 20 h.

Inhibition of cellular growth by gene knockdown was estimated by two methods, colony formation and growth rate measurement. Colony formation was observed by spotting serial dilutions of cell culture on EMM2 plates after CRISPRi induction described above, followed by incubation at 33°C for 3 days. The maximum growth rate was measured as previously described, with some modifications (Makanae et al., 2013). A cell culture after CRISPRi induction was diluted two-fold with EMM2 and was cultured at 33°C for 24 h. To adjust cell density to levels suitable for optical density (OD) measurement, the resulting culture was diluted ten-fold with EMM2, and the diluted culture was further incubated at 33°C for 48 h, while monitoring the OD₆₀₀ every 30 min, using a microplate reader HiTS (SCINICS Co., Ltd., Tokyo, Japan). Growth rate (cell divisions/hour) was calculated as the slope of the line best fit to every nine data points of log₂(OD₆₀₀) versus time (hour), using the least squares method. Maximum growth rate (divisions/hour) is the highest growth rate during the 48 h. The reciprocal of the maximum growth rate indicates minimum doubling time (h).

Transcription of target genes was quantified using reverse transcription quantitative PCR (RT-qPCR), as previously reported (Lyne et al., 2003). In brief, after CRISPRi induction, cells were washed with sterilized water and stored at −80°C until total RNA preparation. Total RNA was prepared by acid phenol-chloroform extraction. cDNA was prepared using a ReverTra Ace qPCR RT kit according to manufacturer instructions (TOYOBO Co., Ltd., Tsuruga, Japan). cDNAs were quantified with a real-time PCR instrument (LightCycler, Roche, Ltd., Basel, Switzerland) and FastStart Essential DNA Green Master Mix (Roche, Ltd., Basel, Switzerland), under standard reaction conditions (pre-incubation at 95°C for 10 min, then 45 cycles of 95°C for 10 s, 60°C for 10 s, and 72°C for 10 s, followed by melting from 65°C to 97°C). Amplification of a single PCR product with each primer set was confirmed by melt curve analysis. mRNA levels of target genes were quantified as values relative to that of act1⁺. Data show the mean±s.d. from at least three biological replicates. Each value of the biological replicates was calculated as the mean of three technical replicates of qPCR measurements. One data point of pSPdCas9 (ns) was excluded from a data analysis shown in the Fig. S2B as an outlier (its value is higher than the mean by two-fold of the standard deviation). All the other data points were included in data analyses. DNA sequences of qPCR primers are shown in Table S2.

Liquid EMM2+T20 medium (2 ml) was inoculated with a single colony grown on an EMM2+T20 plate and incubated at 33°C for 24 h with shaking. This culture was diluted with 10 ml of EMM2+T20 and was grown to log-phase. An aliquot of this log-culture before CRISPRi induction was recovered and fixed for cell morphology analysis. In order to remove thiamine, the other part of the log-culture was washed six times with 1 ml of sterilized water and then resuspended in 1 ml of water. Washed cells were diluted and grown 48 h in EMM2 medium at 33°C with shaking. During this incubation, the cell culture was diluted twice so as not to reach stationary phase. Cells before and after CRISPRi induction were fixed with 2.5% of glutaraldehyde on ice for 30 min. Fixed cells were washed twice with phosphate-buffered saline (PBS) and subjected to fluorescence microcopy in presence of 25 μg/ml DAPI using a fluorescence microscope (EVOS, Thermo Fisher Scientific Inc., Waltham, USA).

Plasmids pSPdCas9, pSPdCas9.2, pSPdCas9.3, pSPdCas9.5, pdSpG.2, pDS_sa2, and pdSpG.5 will be available at National BioResource Project (https://yeast.nig.ac.jp/yeast/top.xhtml). Other strains and plasmids are available upon request.

We thank our colleagues Dr Yusuke Toyoda, Fumie Masuda, Yuri Okabe and Hidefumi Tokunou for valuable discussions.