The authors thank Dr. Gennifer Mager for reading and helpful comments on the manuscript. This work was supported by Ball State University laboratory startup funds and NIH-1R15AI130950-01 to D.A.B.

1. Identification and Cloning of Guide RNA Sequence
<ol>
	<li>Identification of guide RNA sequence
	<ol>
		<li>Identify a 5&#8217;-NGG-3&#8217; PAM sequence close to where the stop codon will be inserted. (Figure 1A) Labeled are all PAM sequences found in the first 100 base pairs of TPK2 (Figure 1A). 
		NOTE: Guide sequences targeting each C. albicans open reading frame can be found at http://osf.io/ARDTX 11,12.</li>
		<li>Identify the Forward Guide Primer_3 sequence, which will be the 20 bases directly upstream of a NGG PAM site and will not contain more than 5 Ts in a row. Left-click on the base directly upstream of the NGG and drag the cursor 20 bases, then left-click on the primer tab to add the primer.</li>
		<li>Identify the Reverse Guide Primer_3 sequence, which will be the complement to the Forward Guide sequence. 
		NOTE: Shown are guides that use PAM_3 (Figure 1B).</li>
		<li>Right-click the primer and select &#8220;copy primer data&#8221;. Paste the sequences into a text editing program.</li>
	</ol>
	</li>
	<li>Add overhang sequences to Forward and Reverse Guide oligos to facilitate cloning (Table 1, Figure 1B).
	<ol>
		<li>Add the nucleotide sequence ATTTG to the 5&#8217; end of the Forward Guide Primer_3 before purchasing.</li>
		<li>Add G to the 3&#8217; end of the Forward Guide Primer_3 before purchasing.</li>
		<li>Add the nucleotide sequence AAAAC to the 5&#8217; end of the Reverse Guide Primer_3 before purchasing.</li>
		<li>Add C to the 3&#8217; end of the Reverse Guide Primer_3 before purchasing.</li>
	</ol>
	</li>
	<li>Digest CaCas9 expression vector pV1524 with BsmBI. 
	NOTE: pV1524 contains an ampicillin (Ampr) and nourseothricin (Natr) markers. Cas9 has been codon-optimized for C. albicans.
	<ol>
		<li>Digest the plasmid by adding: 2 &#956;g of pv1524, 5 &#956;L of 10x Buffer, 1 &#956;L of BsmBI, and H2O to 50 &#956;L in a 1.5 mL tube. Incubate at 55 &#176;C for 20 min. (Alternatively, digest pv1524 for 15 min with Esp3I, an isoschizomer of BsmBI, at 37 &#176;C.)</li>
		<li>Cool to room temperature (RT) and spin for 30 s at 2,348 x g to bring condensation to the bottom of the tube. Proceed to step 1.4 or store the digested plasmid at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Phosphatase-treat the digested backbone.
	<ol>
		<li>Add 1 &#956;L of calf intestinal phosphatase (CIP) to the digestion mixture and incubate at 37 &#176;C for 1 h.</li>
		<li>Purify the digested plasmid using a commercially available polymerase chain reaction (PCR) purification kit (instructions provided with kit) and elute it in 30 &#956;L of elution buffer (EB).</li>
	</ol>
	</li>
	<li>Phosphorylate and anneal Forward Guide Primer_3 and Reverse Guide Primer_3.
	<ol>
		<li>Add 0.5 &#956;L of 100 &#956;M Forward Guide Primer_3, 0.5 &#956;L of 100 &#956;M Reverse Guide Primer_3, 5 &#956;L of 10x T4 ligase buffer, 1 &#956;L of T4 polynucleotide kinase, and 43 &#956;L of H2O to a PCR tube.</li>
		<li>Add 5 &#956;L of 10x T4 ligase buffer, 1 &#956;L of T4 polynucleotide kinase, and 44 &#956;L of molecular biology-grade H2O in a second PCR tube. 
		NOTE: This will serve as the negative control.</li>
		<li>Incubate the reaction mixtures in a thermocycler at 37 &#176;C for 30 min, then at 95 &#176;C for 5 min.</li>
		<li>Cool the mixture at the slowest ramp rate to 16 &#176;C to anneal the ,oligos. Then place the annealed oligo mixture at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Ligate the annealed oligos into digested pv1524 from step 1.4.3.
	<ol>
		<li>Add the following to a PCR tube: 1 &#956;L of 10x T4 ligase buffer, 0.5 &#956;L of T4 DNA ligase, 0.5 &#956;L of annealed oligo mix, digested CIP-treated purified plasmid (20&#8211;40 ng), and H2O to a 10 &#956;L total volume.</li>
		<li>Add the following to a PCR tube: 1 &#956;L of 10x T4 ligase buffer, 0.5 &#956;L of T4 DNA ligase, digested CIP-treated purified vector (20&#8211;40 ng), 1 &#956;L of negative control mixture, and H2O up to a 10 &#956;L total volume.</li>
		<li>Incubate both tubes in a thermocycler at 16 &#176;C for 30 min, then at 65 &#176;C for 10 min, then cool to 25 &#176;C.</li>
	</ol>
	</li>
	<li>Transform 5 &#956;L of the ligation mixtures into chemically competent Escherichia coli DH5&#945; using a standard heat shock transformation protocol. Select on LB Amp/Nat media (200 &#956;g/mL Amp, 50 &#956;g/mL Nat). 
	NOTE: Failure to select pV1524 and its derivatives on double selection media will result in loss of Nat/CaCas9/guide module by FLP/FRT excision in bacteria.</li>
	<li>Purify plasmids from four transformants by miniprep, and sequence the insertion sequence with sequencing primer (Table 1). 
	Note: Most of the time, sequencing 4 transformants is sufficient to identify at least 1 correct clone.</li>
	<li>Save plasmids that have the guide RNA sequence cloned a single time into the BsmBI cut site at -20 &#176;C.</li>
</ol>
2. Designing and Generation of Repair Template
<ol>
	<li>Insert a stop codon by left-clicking in the gene sequence and adding nucleotides that encode a stop codon and restriction digestion site (Figure 1C, Table 1). 
	NOTE: The insertion will disrupt the PAM sequence. 
	NOTE: A restriction digestion site will be included in the repair template sequence to facilitate efficient screening of clones (Figure 1C).</li>
	<li>Left-click 10 bases downstream of where the mutation will be made and drag the cursor 60 bases upstream. Left-click on the primer tab to add the primer. This will add Repair Template Forward_3. Left-click 10 bases upstream of where the mutation will be made and drag the cursor 60 bases downstream. Left-click on the primer tab to add the primer. This will add Repair Template Reverse 3. (Figure 1C)</li>
	<li>Perform primer extension to generate the repair template.
	<ol>
		<li>Add 1.2 &#956;L of 100 &#956;m repair template forward primer, 1.2 &#956;L of 100 &#956;m repair template reverse primer, 6 &#956;L of deoxynucleotide triphosphates (dNTPs) (total concentration 40 mM), 6 &#956;L of buffer, 0.6 &#956;L of Taq polymerase (3 units), and 45 &#956;L of H2O to each of the 4 PCR tubes.</li>
		<li>Perform primer extension by running between 20 and 30 rounds of PCR. Example extension conditions: 2 min at 95 &#176;C, (30 s at 95 &#176;C, 1 min at 50 &#176;C, 1 min at 68 &#176;C) x 34, 10 min at 68 &#176;C.</li>
		<li>Combine contents of all 4 PCR tubes in a 1.5 mL tube and use a PCR purification kit to purify products in 50 &#956;L of H2O.</li>
		<li>Quantitate the primer extension products to ensure sufficient DNA by determining the absorbance at 260 nm. 
		NOTE: Typical final concentration of the primer extension product is ~200&#8211;300 ng/&#181;L.</li>
	</ol>
	</li>
</ol>
3. Transformation of C. albicans with Repair Template and Plasmid
<ol>
	<li>Make TE/lithium acetate.
	<ol>
		<li>Mix 10 mM Tris-Cl, 1 mM EDTA, 100 mM lithium acetate, and H2O (all stock solution pH 7.5) to achieve 50 mL total volume.</li>
	</ol>
	</li>
	<li>Make PLATE
	<ol>
		<li>Mix 40% PEG 3350, 100 mM lithium acetate, 10 mM Tris-Cl, 1 mM EDTA, and H2O (all stock solution pH 7.5) to achieve 50 mL total volume.</li>
	</ol>
	</li>
	<li>Digest correctly-cloned plasmids from step 1.9.
	<ol>
		<li>Add 10 &#956;g of plasmid, 4 &#181;L of 10x buffer, 0.4 &#181;L of 10 mg/mL bovine serum albumin (BSA), 0.5 &#181;L of KpnI, 0.5 &#181;L of SacI, and H20 to 40 &#181;L total volume in a 1.5 mL tube. Incubate at 37 &#176;C overnight (Figure 1D).</li>
	</ol>
	</li>
	<li>Grow an overnight culture of C. albicans SC5314, wild-type prototroph, at 25 &#176;C in yeast peptone dextrose supplemented with 0.27 mM Uridine (YPD + Uri), ideally to OD600 less than 6.
	<ol>
		<li>Pellet 5 OD600 units of cells per transformation by spinning for 5 min at 2,348 x g and suspend the 5 OD600 of pelleted cells in 100 &#181;L TE/lithium acetate.</li>
	</ol>
	</li>
	<li>Add the following to a 1.5 mL tube in the order listed: 1) 100 &#181;L of cells from step 3.4.1, 2) 40 &#181;L of boiled and quick-cooled salmon sperm DNA (10 mg/mL), 3) 10 &#181;g of plasmid digestion from step 3.3.1, 4) 6 &#181;g of purified repair template, and 5) 1 mL of PLATE.</li>
	<li>Add the following to a 1.5 mL tube in the order listed: 1) 100 &#181;L of cells, 2) 40 &#181;L of boiled and quick-cooled salmon sperm DNA (10 mg/mL), 3) H2O volume equal to that of transforming DNA in step 3.5, and 4) 1 mL of PLATE. 
	Note: This will serve as a negative control.</li>
	<li>Mix the transformations gently by pipetting and let incubate at 25 &#176;C overnight.</li>
	<li>Heat shock the cells by placing them in a 44 &#176;C water bath for 25 min.</li>
	<li>Spin for 5 min at 2,348 x g in a benchtop centrifuge and remove the PLATE mixture supernatant. Wash once by adding 1 mL of YPD + Uri and centrifuge again for 5 min at 2,348 x g.</li>
	<li>Suspend the cells in 0.1 mL of YPD + Uri and incubate on a roller drum or shaker at 25 &#176;C overnight.</li>
	<li>Plate on YPD + Uri with 200 &#956;g/mL nourseothricin. Colonies will appear in 2&#8211;5 days.</li>
</ol>
4. Streaks for Single Colonies
<ol>
	<li>Divide a 100 mm x 15 mm Petri dish into quarters, and label each quadrant.
	<ol>
		<li>Touch one of the colonies from the transformation plate with a sterile toothpick or applicator and streak across the longest side of the quadrant.</li>
		<li>Streak for single colonies using an aseptic technique and allow the colonies to grow at 30 &#176;C for 2 days.</li>
	</ol>
	</li>
</ol>
5. Colony PCR
<ol>
	<li>Design the forward check primer (FCP) ~200 base pairs upstream of the restriction site that was introduced and the reverse check primer (RCP) ~300 base pairs downstream.
	<ol>
		<li>Add 0.3 &#956;L of FCP, 0.3 &#956;L of RCP, 0.3 &#956;L of thermostable polymerase (ExTaq 1.5 units), 3 &#956;L of dNTPs (total concentration 40 mM), 3 &#956;L of ExTaq Buffer, and 23 &#956;L of H2O to a 1.5 mL tube. 
		NOTE: Addition of 0.5 &#956;L/reaction dimethyl sulfoxide (DMSO) can improve PCR efficiency.</li>
		<li>Add 0.25 &#956;L of a single yeast colony from step 4.1.2 to the mixture from step 5.1.1, using a P10 pipette tip and taking care not to disturb the agar.</li>
		<li>Amplify DNA by PCR and run 5 &#956;L of the PCR on a gel to ensure amplification is successful, taking care not to disturb the cell pellet at the bottom of the tube.</li>
	</ol>
	</li>
</ol>
6. Restriction Digestion of Colony PCR
<ol>
	<li>Add 10 &#956;L of PCR product (taking care not to not disturb the cell pellet at the bottom of the tube), 3 &#956;L of buffer, 1 &#956;L of restriction enzyme, and 16 &#956;L of H2O, then incubate according to manufacturer&#8217;s instructions and resolve on an agarose gel to identify correct genome edits. 
	NOTE: The restriction enzyme used here is the site encoded in the TPK2 specific repair template.</li>
</ol>
7. Saving Strains
<ol>
	<li>Grow an overnight culture from colonies that have been confirmed by restriction digestion in YPD + Uri at 30 &#176;C.</li>
	<li>Add 1 mL of culture and 1 mL of 50% glycerol (bringing the final concentration of glycerol to 25%) to a tube suitable for storage at -80 &#176;C.</li>
	<li>Store the correct clones at -80 &#176;C. 
	NOTE: Correct strains can be stored at -80 &#176;C for many years.</li>
</ol>
8. Removal of Natr Marker
<ol>
	<li>Streak correct transformant onto yeast peptone maltose (YPMaltose) (2% maltose).</li>
	<li>Pick a colony from the streaked plate and culture the yeast for 48 h at 30 &#176;C in liquid YPMaltose 20 g/L maltose.</li>
	<li>Plate 200&#8211;400 cells on YPMaltose 20 g/L maltose and incubate at 30 &#176;C for 24 h.</li>
	<li>Replicate the plate onto YPMaltose and YPMaltose 200 &#956;g/mL Nat.</li>
	<li>Incubate at 30 &#176;C for 24 h. 
	NOTE: Colonies that no longer grow on YPMaltose 200 &#956;g/mL nourseothricin but grow on YPMaltose have lost the Natr marker (CaCAS9) and guide RNA.</li>
	<li>Save the strains that have lost the Natr marker (CaCAS9) and guide RNA as done in steps 7.1&#8211;7.3. 
	NOTE: A similar plasmid, pV1393, uses the SAP2 as opposed to a MAL2 promotor. Growth on YCB&#8211;BSA will induce flippase and removal of Natr if pV1393 is used for gene editing.</li>
</ol>

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The authors have nothing to disclose.

C. albicans CRISPR efficiently edits the C. albicans genome. pV1524 encodes a yeast codon-optimized Cas9 and is designed such that investigators can easily clone guide RNA sequences downstream of the CaSNR52 promoter (Figure 1)11. It must be ensured that only a single copy of the guide sequence has been cloned into CaCas9 expression vectors by sequencing, as extra copies will impede genome editing. If multiple copies of the guide are introduced consistently, one should lower the concentration of the annealed guide used in ligation. The vector and protocol described allow targeting of any C. albicans gene. Although C. albicans is diploid, only a single transformation is required to target both alleles of a gene. Furthermore, the processive nature of CRISPR-CaCas9 genome editing enables researchers to target multiple members of gene families. Many gene families such as the secreted aspartyl proteases (SAPS) and agglutinin-like sequence proteins (ALS) are important for C. albicans virulence. CRISPR genome editing will facilitate investigation of these gene families.
The protocols described above introduce a stop codon to an open reading frame, resulting in the phenotypic equivalent of a null (Figure 2). A wide variety of genetic alternations can be made by varying the repair template. Nonsense, missense, and silent mutations can be inserted via recombination with an appropriate repair template. Incorporation of a restriction site streamlines transformant screening, as those without must be screened by sequencing12,13. In addition, C. albicans CRISPR enables researchers to generate insertions and deletions, making it an ideal system to insert affinity tags, perform promotor swaps, and generate knockouts (Figure 3). Screening for correct transformants for these mutations is more laborious, as it is necessary to sequence the edits to confirm correct incorporation of the repair templates. Furthermore, Southern blot may be necessary to ensure additional copies of a gene have not been inserted at additional locations in the genome. The requirement of the NGG PAM site places slight limitations on the regions of the genome that can be targeted. The development of alternative CRISPR systems that use alternative nucleases such as Cpf1 or variations on the Cas9 system have/will alleviate many of these limitations14. To the investigators&#39; knowledge at this time, these systems have not yet been applied to C. albicans.
The CRISPR system described in the above protocol has been developed such that it can be applied in a wide variety of species including Saccharomyces cerevisiae, Naumouozyma castellii, and the human pathogen Candida glabrata11. Transformation and efficient editing of these yeast requires slight changes to the described protocol, but the framework for editing these alternate genomes is remarkably similar to that described for C. albicans12. Furthermore, yeast provide an excellent mechanism to develop genome editing procedures. In yeast, when ADE2 is mutated, a precursor to the adenine biosynthesis pathway accumulates, turning the cells red. This easily observable phenotype allows investigators to identify edited cells and quickly troubleshoot genome editing protocols. Combined with the extensive molecular biology toolbox available for fungi, protocols for editing numerous yeast species have been developed15,16. Such a broad application of genome editing technology in fungi has the potential to significantly impact a wide variety of scientific disciplines.
CRISPR has greatly improved the efficiency of genome engineering in C. albicans, but to date CRISPR has not been used to perform genome wide screens in C. albicans. Current protocols require a repair template to introduce mutations, as the nonhomologous end joining pathway in C. albicans is inefficient12. The generation of repair template oligos for every gene is a significant barrier to the execution of genome-wide screens. The confluence of decreased costs of DNA synthesis and advances to CRISPR technologies will make development of deletion libraries more feasible. For instance, expression of a repair template from the CaCas9 vector paves the way for the development of sustainable plasmid libraries that target every gene11. Furthermore, transient Candida CRISPR protocols that do not require CaCas9 expression vector incorporate into the C. albicans genome have been developed17. In addition, increased guide expression increases genome editing efficiency18. These, and other advances to CRISPR technologies, are crucial to the development of genome-wide screens in C. albicans19,20,21,22.
The C. albicans genome is diploid, but A and B alleles are not always identical5. Such heterozygosity provides both challenges and opportunities. If one aims to target both alleles, a PAM site, guide sequence, and repair template that will act on both copies of the gene must be used. However, depending upon single nucleotide polymorphisms present in a gene, the C.albicans CRISPR system enables investigators to target a single allele. Such precision has the potential to allow investigators to examine functional differences between alleles. Targeting specific alleles must be done carefully, as loss of heterozygosity (LOH) at an allele or of an entire chromosome has been observed. When editing single C. albicans alleles, one must examine adjacent DNA sequences to determine if a clone has maintained a diploid SNP profile. In addition, off-target effects are quite low for C. albicans CRISPR, but whole genome sequencing can be considered for key strains.

Candida albicans is the most prevalent human fungal pathogen1,2,3. Understanding differences between C. albicans and mammalian molecular biology is critical to development of the next generation of antifungal therapeutics. This requires investigators to be able to quickly and accurately genetically manipulate C. albicans.
Genetic manipulation of C. albicans has historically been challenging. C. albicans does not maintain plasmids, thus all constructs must be incorporated into the genome. Furthermore, C. albicans is diploid; therefore, when knocking out a gene or introducing a mutation, it is important to ensure that both copies have been changed4. In addition, some C. albicans loci are heterozygous, further complicating genetic interrogation5. To genetically manipulate C. albicans, it is typical to perform multiple rounds of homologous recombination6. However, the diploid nature of the genome and laborious construct development have made this a potentially tedious process, especially if multiple changes are required. These limitations and the medical importance of C. albicans demand the development of new technologies that enable investigators to more easily manipulate the C. albicans genome.
Clustered regularly interspaced short palindromic repeats (CRISPR)-mediated genome editing is a powerful tool that allows researchers to change the sequence of a genome. CRISPR requires three components: 1) the Cas9 nuclease that cleaves the target DNA, 2) a 20 base guide RNA that targets Cas9 to the sequence of interest, and 3) repair template DNA that repairs the cleavage site and incorporates the intended change7,8. Once the guide brings Cas9 to the target genome sequence, Cas9 requires a protospacer adjacent motif (PAM) sequence (NGG) directly upstream of the guide sequence to cleave the DNA9. The requirement for both the 20 base guide and PAM sequences provides a high degree of targeting specificity and limits off-target cleavage.
CRISPR systems have been designed to edit the genomes of a diverse set of organisms and tackle a wide variety of problems10. Described here is a flexible, efficient CRISPR protocol for editing a C. albicans gene of interest. The experiment introduces a stop codon to a gene, causing translation termination. Other edits can be made depending on the repair template that is introduced. A fragment marked with nourseothricin (Natr) containing yeast codon-optimized Cas9 (CaCas9) and a guide RNA is incorporated into the C. albicans genome at a neutral site. Cotransformation with the repair template encoding the desired mutation leads to repair of the cleavage by homologous recombination and efficient genome editing. Described below is the editing of TPK2, but all C. albicans open reading frames can be targeted multiple times by CRISPR. The CRISPR system is flanked by FRT sites and can be removed from the C. albicans genome by induction of flippase encoded on the CaCas9 expression plasmid. The C. albicans CRISPR system enables investigators to accurately and quickly edit the C. albicans genome11,12.

<table><tbody><tr><td>&lt;strong&gt;Chemicals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Agar</td><td>BD Bacto</td><td>214010</td><td></td></tr><tr><td>agarose</td><td>amresco</td><td>0710-500G</td><td></td></tr><tr><td>Ampicillan</td><td>Sigma-Aldrich</td><td>A9518</td><td></td></tr><tr><td>Bacto Peptone</td><td>BD Bacto</td><td>211677</td><td></td></tr><tr><td>Bsmb1</td><td>New England Biolabs</td><td>R0580L</td><td></td></tr><tr><td>Calf intestinal phosphatase (CIP)</td><td>New England Biolabs</td><td>M0290L</td><td></td></tr><tr><td>Cut Smart Buffer</td><td>New England Biolabs</td><td>B7204S</td><td></td></tr><tr><td>Dimethyl Sulfoxide (DMSO)</td><td>Sigma-Aldrich</td><td>D8418</td><td></td></tr><tr><td>dNTPs</td><td>New England Biolabs</td><td>N0447L</td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Sigma-Aldrich</td><td>3609</td><td></td></tr><tr><td>Glacial Acetic Acid</td><td>Sigma-Aldrich</td><td>2810000ACS</td><td></td></tr><tr><td>Glucose</td><td>BDH VWR analytical</td><td>BDH9230</td><td></td></tr><tr><td>Glycerol</td><td>Sigma-Aldrich</td><td>49767</td><td></td></tr><tr><td>Kpn1</td><td>New England Biolabs</td><td>R3142L</td><td></td></tr><tr><td>LB-Medium</td><td>MP</td><td>3002-032</td><td></td></tr><tr><td>Lithium Acetate</td><td>Sigma-Aldrich</td><td>517992</td><td></td></tr><tr><td>L-Tryptophan</td><td>Sigma-Aldrich</td><td>T0254</td><td></td></tr><tr><td>Maltose</td><td>Sigma-Aldrich</td><td>M5885</td><td></td></tr><tr><td>Molecular Biology Water</td><td>Sigma-Aldrich</td><td>W4502</td><td></td></tr><tr><td>NEB3.1 Buffer</td><td>New England Biolabs</td><td>B7203S</td><td></td></tr><tr><td>Nourseothricin</td><td>Werner Bioagents</td><td>74667</td><td></td></tr><tr><td>Poly(ethylene glycol) PEG 3350</td><td>Sigma-Aldrich</td><td>P4338</td><td></td></tr><tr><td>Sac1</td><td>New England Biolabs</td><td>R3156L</td><td></td></tr><tr><td>Salmon Sperm DNA</td><td>Invitrogen</td><td>AM9680</td><td></td></tr><tr><td>T4 Polynucleotide kinase</td><td>New England Biolabs</td><td>M0201S</td><td></td></tr><tr><td>T4 DNA ligase</td><td>New England Biolabs</td><td>M0202L</td><td></td></tr><tr><td>Taq polymerase</td><td>New England Biolabs</td><td>M0267X</td><td></td></tr><tr><td>Tris HCl</td><td>Sigma-Aldrich</td><td>T3253</td><td></td></tr><tr><td>uridine</td><td>Sigma-Aldrich</td><td>U3750</td><td></td></tr><tr><td>Yeast Extract</td><td>BD Bacto</td><td>212750</td><td></td></tr><tr><td>&lt;strong&gt;Equipment &lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Electrophoresis Appartus</td><td></td><td></td><td></td></tr><tr><td>Incubator</td><td></td><td></td><td></td></tr><tr><td>Microcentifuge</td><td></td><td></td><td></td></tr><tr><td>PCR machine</td><td></td><td></td><td></td></tr><tr><td>Replica Plating Apparatus</td><td></td><td></td><td></td></tr><tr><td>Rollerdrum or shaker</td><td></td><td></td><td></td></tr><tr><td>Spectrophotometer</td><td></td><td></td><td></td></tr><tr><td>Waterbath</td><td></td><td></td><td></td></tr></tbody></table>

crispr-mediated genome editing of the human fungal pathogen candida albicans

This method describes the efficient CRISPR mediated genome editing of the diploid human fungal pathogen Candida albicans. CRISPR-mediated genome editing in C. albicans requires Cas9, guide RNA, and repair template. A plasmid expressing a yeast codon optimized Cas9 (CaCas9) has been generated. Guide sequences directly upstream from a PAM site (NGG) are cloned into the Cas9 expression vector. A repair template is then made by primer extension in vitro. Cotransformation of the repair template and vector into C. albicans leads to genome editing. Depending on the repair template used, the investigator can introduce nucleotide changes, insertions, or deletions. As C. albicans is a diploid, mutations are made in both alleles of a gene, provided that the A and B alleles do not harbor SNPs that interfere with guide targeting or repair template incorporation. Multimember gene families can be edited in parallel if suitable conserved sequences exist in all family members. The C. albicans CRISPR system described is flanked by FRT sites and encodes flippase. Upon induction of flippase, the antibiotic marker (CaCas9) and guide RNA are removed from the genome. This allows the investigator to perform subsequent edits to the genome. C. albicans CRISPR is a powerful fungal genetic engineering tool, and minor alterations to the described protocols permit the modification of other fungal species including C. glabrata, N. castellii, and S. cerevisiae.

Sequences of guide RNAs and repair templates that target C. albicans TPK2, a c-AMP kinase catalytic subunit, were designed according to the guidelines suggested above. Sequences are shown in (Table 1, Figure 1). Guide RNAs were cloned into CaCas9 expression vectors and cotransformed with repair template in wild-type C. albicans. An EcoRI restriction digestion site and stop codons in the repair template disrupt the PAM site and facilitate screening for correct mutants (Figure 1). Transformants were streaked for single colonies and screened by colony PCR and restriction digestion for incorporation of the repair template (Figure 2). Restriction digestion quickly distinguishes wild-type from mutant sequences.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Oligonucleotide Name </td>
			<td>Oligonucleotide Sequence </td>
		</tr>
		<tr>
			<td>Forward Guide Primer_3</td>
			<td>ATTTGgggtgaactatttgttcgccG</td>
		</tr>
		<tr>
			<td>Reverse Guide Primer_3</td>
			<td>AAAACggcgaacaaatagttcacccC</td>
		</tr>
		<tr>
			<td>Repair Template Forward_3</td>
			<td>tcagcaatatcagcaacaatttcaacaaccgcagcaacaactttatTAAGAATTCggcga</td>
		</tr>
		<tr>
			<td>Repair Template Reverse_3</td>
			<td>attttgtccagtttgggctgcagcagggtgaactatttgttcgccGAATTCTTAataaag</td>
		</tr>
		<tr>
			<td>Forward Check Primer</td>
			<td>ttaaagaaacttcacatcaccaag</td>
		</tr>
		<tr>
			<td>Reverse Check Primer</td>
			<td>actttgatagcataatatctaccat</td>
		</tr>
		<tr>
			<td>Sequencing Primer</td>
			<td>ggcatagctgaaacttcggccc</td>
		</tr>
	</tbody>
</table>
Table 1: List of oligo nucleotides used for this study. Sites added for cloning purposes are capitalized and bolded in the guide primer sequences. Sequences in the repair template that mutate the genomic DNA are capitalized and bolded.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58764/58764fig1.jpg" /> 
Figure 1: Diagram of guide RNA and repair template design. (A) Labeling of all the PAM sequences in the first 100 nucleotides of TPK2. PAM sequence 3 (PAM_3) is highlighted, as that is the sequence used in this study. (B) Guide RNA design using PAM_3. 20 base primers designed using SnapGene are lowercase and blue. Additional bases required for cloning are uppercase and green shown offset. (C) Repair template primers that insert a TAA stop codon and EcoRI site are inserted into the TPK2 reading frame. DNA that differs from the wild-type sequence is red and uppercase. (D) Example of how a guide is designed on the positive strand of DNA using PAM_4. (E) Schematic diagram of pV1524 after cloning of the guide RNA and digestion with KpnI and SacI. Neut5L-5&#39; and Neut5L-3&#39; target the vector to the Neut5L site in the C. albicans genome. CaENO1p is the promotor that drives expression of the yeast-optimized CaCas9. Natr is the nourseothricin resistance cassette. CaSNR52p is the promotor driving guide RNA expression (sgRNA). FRT sites are cleaved and recombined by flippase (FLP) removing the CRISPR cassette upon flippase expression. A schematic similar to (E) was published by Vyas et al.11. <a href="https://www.jove.com/files/ftp_upload/58764/58764fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58764/58764fig2.jpg" /> 
Figure 2: Introduction and confirmation of a stop codon and EcoRI restriction site to TPK2. Primers used for amplification are listed in Table 1. Wild-type and mutant sequences are shown below the gel. This figure has been modified from Vyas et al.11. <a href="https://www.jove.com/files/ftp_upload/58764/58764fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58764/58764fig3.jpg" /> 
Figure 3: Cartoon description of repair templates that will generate (A) deletions and (B) insertions. Grey dashes in (A) depict intervening sequences not present in the repair template primers. <a href="https://www.jove.com/files/ftp_upload/58764/58764fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about CRISPR-mediated Genome Editing of the Human Fungal Pathogen Candida albicans at JoVE.com

CRISPR-mediated Genome Editing of the Human Fungal Pathogen Candida albicans

Efficient genome engineering of Candida albicans is critical to understanding the pathogenesis and development of therapeutics. Here, we described a protocol to quickly and accurately edit the C. albicans genome using CRISPR. The protocol allows investigators to introduce a wide variety of genetic modifications including point mutations, insertions, and deletions.

CRISPR-mediated Genome Editing of the Human Fungal Pathogen Candida albicans

This method describes the efficient CRISPR mediated genome editing of the diploid human fungal pathogen Candida albicans. CRISPR-mediated genome editing ...

crispr-mediated-genome-editing-human-fungal-pathogen-candida

Research

JoVE Journal

Genetics

11.7K Views.  Ball State University. Deze methode kan helpen onthullen de verschillen tussen schimmels en zoogdieren op moleculair niveau. Dergelijke verschillen kunnen worden benut om nieuwe therapeutische benaderingen te identificeren. Het belangrijkste voordeel van deze techniek is dat het het genoom van C.Albicans efficiënter wijzigt dan klassieke homologe recombinatietechnieken.Om de RNA-reeks voor hulpgidsen te identificeren, identificeert u een NGG PAM-reeks in de buurt van waar de stopcodon wordt ingevoegd. Hier ...

Video: CRISPR-mediated Genome Editing of the Human Fungal Pathogen Candida albicans

Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

Gene deletion plays an important role in the analysis of gene function. One of the most efficient methods to disrupt genes in a targeted manner is the replacement of the entire gene with a selectable marker via homologous recombination. During homologous recombination, exchange of DNA takes place between sequences with high similarity. Therefore, linear genomic sequences flanking a target gene can be used to specifically direct a selectable marker to the desired integration site. Blunt ends of the deletion construct activate the cell&#39;s DNA repair systems and thereby promote integration of the construct either via homologous recombination or by non-homologous-end-joining. In organisms with efficient homologous recombination, the rate of successful gene deletion can reach more than 50% making this strategy a valuable gene disruption system. The smut fungus Ustilago maydis is a eukaryotic model microorganism showing such efficient homologous recombination. Out of its about 6,900 genes, many have been functionally characterized with the help of deletion mutants, and repeated failure of gene replacement attempts points at essential function of the gene. Subsequent characterization of the gene function by tagging with fluorescent markers or mutations of predicted domains also relies on DNA exchange via homologous recombination. Here, we present the U. maydis strain generation strategy in detail using the simplest example, the gene deletion.

Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

We describe a robust gene replacement strategy to genetically manipulate the smut fungus Ustilago maydis. This protocol explains how to generate deletion mutants to investigate infection phenotypes. It can be extended to modify genes in any desired way, e.g., by adding a sequence encoding a fluorescent protein tag.

Gene deletion plays an important role in the analysis of gene function. One of the most efficient methods to disrupt genes in a targeted manner is the ...

Generation of Fluorescent Protein Fusions in Candida Species

Candida species, prevalent colonizers of the intestinal and genitourinary tracts, are the cause of the majority of invasive fungal infections in humans. Thus, molecular and genetic tools are needed to facilitate the study of their pathogenesis mechanisms. PCR-mediated gene modification is a straightforward and quick approach to generate epitope-tagged proteins to facilitate their detection. In particular, fluorescent protein (FP) fusions are powerful tools that allow visualization and quantitation of both yeast cells and proteins by fluorescence microscopy and immunoblotting, respectively. Plasmids containing FP encoding sequences, along with nutritional marker genes that facilitate the transformation of Candida species, have been generated for the purpose of FP construction and expression in Candida. Herein, we present a strategy for constructing a FP fusion in a Candida species. Plasmids containing the nourseothricin resistance transformation marker gene (NAT1) along with sequences for either green, yellow, or cherry FPs (GFP, YFP, mCherry) are used along with primers that include gene-specific sequences in a polymerase chain reaction (PCR) to generate a FP cassette. This gene-specific cassette has the ability to integrate into the 3'-end of the corresponding gene locus via homologous recombination. Successful in-frame fusion of the FP sequence into the gene locus of interest is verified genetically, followed by analysis of fusion protein expression by microscopy and/or immuno-detection methods. In addition, for the case of highly expressed proteins, successful fusions can be screened for primarily by fluorescence imaging techniques.

Generation of Fluorescent Protein Fusions in Candida Species

PCR-mediated gene modification can be used to generate fluorescent protein fusions in Candida species, which facilitates visualization and quantitation of yeast cells and proteins. Herein, we present a strategy for constructing a fluorescent protein fusion (Eno1-FP) in Candida parapsilosis.

Candida species, prevalent colonizers of the intestinal and genitourinary tracts, are the cause of the majority of invasive fungal infections in humans. ...

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide RNA (sgRNA) that confers specificity, the Cas9 protein cleaves both DNA strands at the targeted locus. The DNA break can trigger either non-homologous end joining (NHEJ) or homology directed repair (HDR). NHEJ can introduce small deletions or insertions which lead to frame-shift mutations, while HDR allows for larger and more precise perturbations. Here, we present protocols for generating knockout cell lines by coupling established CRISPR/Cas9 methods with two options for downstream selection/screening. The NHEJ approach uses a single sgRNA cut site and selection-independent screening, where protein production is assessed by dot immunoblot in a high-throughput manner. The HDR approach uses two sgRNA cut sites that span the gene of interest. Together with a provided HDR template, this method can achieve deletion of tens of kb, aided by the inserted selectable resistance marker. The appropriate applications and advantages of each method are discussed.

Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide ...

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been co-opted as a powerful tool for precise eukaryotic genome engineering. Here, we present a rapid and simple method using chimeric single guide RNAs (sgRNA) and CRISPR-Cas9 Ribonucleoproteins (RNPs) for the efficient and precise generation of genomic point mutations in C. elegans. We describe a pipeline for sgRNA target selection, homology-directed repair (HDR) template design, CRISPR-Cas9-RNP complexing and delivery, and a genotyping strategy that enables the robust and rapid identification of correctly edited animals. Our approach not only permits the facile generation and identification of desired genomic point mutant animals, but also facilitates the detection of other complex indel alleles in approximately 4 - 5 days with high efficiency and a reduced screening workload.

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Here, we present a method to engineer the genome of C. elegans using CRISPR-Cas9 ribonucleoproteins and homology dependent repair templates.

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been ...

Generation of Defined Genomic Modifications Using CRISPR-CAS9 in Human Pluripotent Stem Cells

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise heterozygous genetic modifications is challenging due to CRISPR-CAS9 mediated indel formation in the second allele. Here, we demonstrate a protocol to help overcome this difficulty by using two repair templates in which only one expresses the desired sequence change, while both templates contain silent mutations to prevent re-cutting and indel formation. This methodology is most advantageous for gene editing coding regions of DNA to generate isogenic control and mutant human stem cell lines for studying human disease and biology. In addition, optimization of transfection and screening methodologies have been performed to reduce labor and cost of a gene editing experiment. Overall, this protocol is widely applicable to many genome editing projects utilizing the human pluripotent stem cell model.

This protocol provides a method to facilitate the generation of defined heterozygous or homozygous nucleotide changes using CRISPR-CAS9 in human pluripotent stem cells.

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise ...

CRISPR-Cas9-Mediated Genome Editing in the Filamentous Ascomycete Huntiella omanensis

The CRISPR-Cas9 genome editing system is a molecular tool that can be used to introduce precise changes into the genomes of model and non-model species alike. This technology can be used for a variety of genome editing approaches, from gene knockouts and knockins to more specific changes like the introduction of a few nucleotides at a targeted location. Genome editing can be used for a multitude of applications, including the partial functional characterization of genes, the production of transgenic organisms and the development of diagnostic tools. Compared to previously available gene editing strategies, the CRISPR-Cas9 system has been shown to be easy to establish in new species and boasts high efficiency and specificity. The primary reason for this is that the editing tool uses an RNA molecule to target the gene or sequence of interest, making target molecule design straightforward, given that standard base pairing rules can be exploited. Similar to other genome editing systems, CRISPR-Cas9-based methods also require efficient and effective transformation protocols as well as access to good quality sequence data for the design of the targeting RNA and DNA molecules. Since the introduction of this system in 2013, it has been used to genetically engineer a variety of model species, including Saccharomyces cerevisiae, Arabidopsis thaliana, Drosophila melanogaster and Mus musculus. Subsequently, researchers working on non-model species have taken advantage of the system and used it for the study of genes involved in processes as diverse as secondary metabolism in fungi, nematode growth and disease resistance in plants, among many others. This protocol detailed below describes the use of the CRISPR-Cas9 genome editing protocol for the truncation of a gene involved in the sexual cycle of Huntiella omanensis, a filamentous ascomycete fungus belonging to the Ceratocystidaceae family.

CRISPR-Cas9-Mediated Genome Editing in the Filamentous Ascomycete Huntiella omanensis

The CRISPR-Cas9 genome editing system is an easy-to-use genome editor that has been used in model and non-model species. Here we present a protein-based version of this system that was used to introduce a premature stop codon into a mating gene of a non-model filamentous ascomycete fungus.

The CRISPR-Cas9 genome editing system is a molecular tool that can be used to introduce precise changes into the genomes of model and non-model species ...

CRISPR/Cas12a Multiplex Genome Editing of Saccharomyces cerevisiae and the Creation of Yeast Pixel Art

High efficiency, ease of use and versatility of the clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9 (CRISPR/Cas9) system has facilitated advanced genetic modification of Saccharomyces cerevisiae, a model organism and workhorse in industrial biotechnology. CRISPR-associated protein 12a (Cas12a), an RNA-guided endonuclease with features distinguishable from Cas9 is applied in this work, further extending the molecular toolbox for genome editing purposes. A benefit of the CRISPR/Cas12a system is that it can be used in multiplex genome editing with multiple guide RNAs expressed from a single transcriptional unit (single CRISPR RNA (crRNA) array). We present a protocol for multiplex integration of multiple heterologous genes into independent loci of the S. cerevisiae genome using the CRISPR/Cas12a system with multiple crRNAs expressed from a single crRNA array construct. The proposed method exploits the ability of S. cerevisiae to perform in vivo recombination of DNA fragments to assemble the single crRNA array into a plasmid that can be used for transformant selection, as well as the assembly of donor DNA sequences that integrate into the genome at intended positions. Cas12a is pre-expressed constitutively, facilitating cleavage of the S.&#160;cerevisiae genome at the intended positions upon expression of the single crRNA array. The protocol includes the design and construction of a single crRNA array and donor DNA expression cassettes, and exploits an integration approach making use of unique 50-bp DNA connectors sequences and separate integration flank DNA sequences, which simplifies experimental design through standardization and modularization and extends the range of applications. Finally, we demonstrate a straightforward technique for creating yeast pixel art with an acoustic liquid handler using differently colored carotenoid producing yeast strains that were constructed.

CRISPR/Cas12a Multiplex Genome Editing of Saccharomyces cerevisiae and the Creation of Yeast Pixel Art

The CRISPR/Cas12a system in combination with a single crRNA array enables efficient multiplex editing of the S.&#160;cerevisiae genome at multiple loci simultaneously. This is demonstrated by constructing carotenoid producing yeast strains which are subsequently used to create yeast pixel art.

High efficiency, ease of use and versatility of the clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9 (CRISPR/Cas9) ...

Bioengineering

Bio-energetics Investigation of Candida albicans Using Real-time Extracellular Flux Analysis

Mitochondria are essential organelles for the cellular metabolism and survival. A variety of key events take place in mitochondria, such as cellular respiration, oxidative metabolism, signal transduction, and apoptosis. Consequently, mitochondrial dysfunction is reported to play an important role in the antifungal drug tolerance and virulence of pathogenic fungi. Recent data have also led to the recognition of the importance of the mitochondria as an important contributor to fungal pathogenesis. Despite the importance of the mitochondria in fungal biology, standardized methods to understand its function are poorly developed. Here, we present a procedure to study the basal oxygen consumption rate (OCR), a measure of mitochondrial respiration, and extracellular acidification rates (ECAR), a measure of glycolytic function in C. albicans strains. The method described herein can be applied to any Candidaspp. strains without the need to purify mitochondria from the intact fungal cells. Furthermore, this protocol can also be customized to screen for inhibitors of mitochondrial function in C. albicans strains.

Bio-energetics Investigation of Candida albicans Using Real-time Extracellular Flux Analysis

Here, we present a stepwise protocol to investigate the mitochondrial respiration and glycolytic function in Candida Albicans using an extra flux analyzer.

Mitochondria are essential organelles for the cellular metabolism and survival. A variety of key events take place in mitochondria, such as cellular ...

Medicine

Use of In Vivo Imaging to Screen for Morphogenesis Phenotypes in Candida albicans Mutant Strains During Active Infection in a Mammalian Host

Candida albicans is an important human pathogen. Its ability to switch between morphologic forms is central to&#160;its pathogenesis; these morphologic changes are regulated by a complex signaling network controlled in response to environmental stimuli. These regulatory components have been highly studied, but almost all studies use a variety of in vitro stimuli to trigger filamentation. To determine how morphogenesis is regulated during the pathogenesis process, we developed an in vivo microscopy system to obtain high spatial resolution images of organisms undergoing hyphal formation within the mammalian host. The protocol presented here describes the use of this system to screen small collections of C. albicans mutant strains, allowing us to identify key regulators of morphogenesis as it occurs at the site of infection. Representative results are presented, demonstrating that some regulators of morphogenesis, such as the transcriptional regulator Efg1, have consistent phenotypes in vitro and in vivo, whereas other regulators, such as adenyl cyclase (Cyr1), have significantly different phenotypes in vivo compared to in vitro.

Use of In Vivo Imaging to Screen for Morphogenesis Phenotypes in Candida albicans Mutant Strains During Active Infection in a Mammalian Host

This manuscript describes a method for screening moderately sized Candida albicans mutant libraries for morphogenesis phenotypes during active infection in a mammalian host using non-invasive confocal microscopy.

Candida albicans is an important human pathogen. Its ability to switch between morphologic forms is central to&#160;its pathogenesis; these morphologic ...

Immunology and Infection

Small-Scale Plasma Membrane Preparation for the Analysis of Candida albicans Cdr1-mGFPHis

The successful biochemical and biophysical characterization of ABC transporters depends heavily on the choice of the heterologous expression system. Over the past two decades, we have developed a yeast membrane protein expression platform that has been used to study many important fungal membrane proteins. The expression host Saccharomyces cerevisiae AD&#916;&#916; is deleted in seven major endogenous ABC transporters and it contains the transcription factor Pdr1-3 with a gain-of-function mutation that enables the constitutive overexpression of heterologous membrane protein genes stably integrated as single copies at the genomic PDR5 locus. The creation of versatile plasmid vectors and the optimization of one-step cloning strategies enables the rapid and accurate cloning, mutagenesis, and expression of heterologous ABC transporters. Here, we describe the development and use of a novel protease-cleavable mGFPHis double tag (i.e., the monomeric yeast enhanced green fluorescent protein yEGFP3 fused to a six-histidine affinity purification tag) that was designed to avoid possible interference of the tag with the protein of interest and to increase the binding efficiency of the His tag to nickel-affinity resins. The fusion of mGFPHis to the membrane protein ORF (open reading frame) enables easy quantification of the protein by inspection of polyacrylamide gels and detection of degradation products retaining the mGFPHis tag. We demonstrate how this feature facilitates detergent screening for membrane protein solubilization. A protocol for the efficient, fast, and reliable isolation of the small-scale plasma membrane preparations of the C-terminally tagged Candida albicans multidrug efflux transporter Cdr1 overexpressed in S. cerevisiae AD&#916;&#916;, is presented. This small-scale plasma membrane isolation protocol generates high-quality plasma membranes within a single working day. The plasma membrane preparations can be used to determine the enzyme activities of Cdr1 and Cdr1 mutant variants.

Small-Scale Plasma Membrane Preparation for the Analysis of Candida albicans Cdr1-mGFPHis

This article presents a small-scale plasma membrane isolation protocol for the characterization of Candida albicans ABC (ATP-binding cassette) protein Cdr1, overexpressed in Saccharomyces cerevisiae. A protease-cleavable C-terminal mGFPHis double tag with a 16-residue linker between Cdr1 and the tag was designed to facilitate the purification and detergent-screening of Cdr1.

The successful biochemical and biophysical characterization of ABC transporters depends heavily on the choice of the heterologous expression system. Over ...

CRISPR-mediated Genome Editing of the Human Fungal Pathogen Candida albicans

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