An Efficient Workflow for Screening and Stabilizing CRISPR/Cas9-Mediated Mutant Lines in Bombyx mori

The domestic silkworm Bombyx mori is extensively studied as a model organism for lepidopteran genetics and has an economic value in silk production. Silkworms also have applications in biomedical and cosmetic industries, and the production of mutant B. mori strains significantly enhances basic and applied silkworm research. In recent years, CRISPR/Cas9 technology is being rapidly adopted as the most efficient molecular tool for generating silkworm lines carrying mutations in target genes. Here we illustrate a complete and efficient workflow to screen, characterize rapidly and follow mutations through generations, allowing the generation of B. mori lines, stably inheriting single CRISPR/Cas9-induced mutations. This approach relies on the use of different molecular methods, the heteroduplex assay, cloning followed by Sanger sequencing, and the amplification refractory mutation system PCR. The use of these methodologies in a sequential combination allows the identification of CRISPR/Cas9-induced mutations in genes mapping on both autosomes and sex chromosomes, and the selection of appropriate individuals to found stable mutant B. mori lines. This protocol could be further applied to screen CRISPR/Cas9 mutations in haploid insects.


Introduction
The domesticated silkworm Bombyx mori has been extensively studied as a model organism, allowing the characterization of numerous biological processes [1,2], and for its economic value in silk production. In addition to the textile industry, silk fiber is also used in the biomedical and cosmetic sectors, and several studies have suggested silkworm larvae are suitable bioreactors, producing valuable recombinant proteins or modified silk useful for applied research [3]. Therefore, the production of modified or mutant B. mori strains has been fundamental for both basic research and applied purposes. Since Tamura and colleagues introduced the transposon-mediated germline transformation in B. mori [4], several techniques, including Zinc Finger Nucleases [5] and Transcription Activator-Like Effector Nucleases (TALENS) [6], have been developed to generate silkworm strains, carrying specific mutations in endogenous genes, or expressing exogenous factors.
In recent years, the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9 has become the dominant method of targeted mutagenesis due to its reduced costs, simplified workflow, and significantly increased efficiency [7]. The mechanism of CRISPR/Cas9 requires a guide RNA (gRNA) and Cas9 endonuclease. Briefly, the gRNA and Cas9 localize to the genomic site of interest, where Cas9 causes a double-strand break in the DNA upstream to a protospacer adjacent motif (PAM). During the subsequent DNA repair by non-homologous end-joining, random mutations frequently occur upstream of the PAM site [7]. These modifications are often indels, ranging from one to tens of base pairs (bp) that can generate null alleles, e.g., carrying premature stop codons in the coding region. Dual injections with two appropriate gRNAs have also been used to delete large target region ( Figure 1). Next, the PCR amplicon is used to confirm that the gRNA and Cas9 efficiently localize to and digest the DNA, in vitro. Once gRNA efficacy has been confirmed, microinjections into fertile B. mori eggs are performed to generate the G0 injected moths ( Figure 2).  A workflow for the initial identification of mutations. (a) G0 injected eggs are reared following microinjection and crossed with wildtype moths to generate (b) G1 egg batches. (c) DNA extraction and PCR are performed on eggs from G1 egg batches followed by (d) temperature denaturing and heteroduplex formation. (e) A PAGE-based heteroduplex assay is performed to identify which egg batches contain mutations. In this scheme G0_1 and G0_3 moths carry mutations in germline cells and give rise to mutant G1 progeny. G0_2 moth carries somatic mutations which are not transmitted to the G1 progeny. G0_4 lacks any mutation.
Molecular screening for the establishment of a stable homozygous gene knockout line involves three primary steps; (1) initial mutant screening with heteroduplex assay, (2) mutation sequence determination with TA cloning and Sanger sequencing, and (3) specific mutation screening by ARMS-PCR. Some authors begin initial screening on G0 injected larvae; however, these animals may be mosaics without the potential for germline transmission of the observed DNA modification. Furthermore, tissue extraction from B. mori larvae is an invasive procedure that can result in mortality. Therefore, we rear all G0 larvae without interference and begin molecular screening on G1 egg batches.

Initial Identification of Mutations
The initial G1 embryo screening for mutations ( Figure 2) involves a heteroduplex assay, which allows for the identification of the egg batch/es carrying promising mutations target region ( Figure 1). Next, the PCR amplicon is used to confirm that the gRNA and Cas9 efficiently localize to and digest the DNA, in vitro. Once gRNA efficacy has been confirmed, microinjections into fertile B. mori eggs are performed to generate the G0 injected moths ( Figure 2).  Molecular screening for the establishment of a stable homozygous gene knockout line involves three primary steps; (1) initial mutant screening with heteroduplex assay, (2) mutation sequence determination with TA cloning and Sanger sequencing, and (3) specific mutation screening by ARMS-PCR. Some authors begin initial screening on G0 injected larvae; however, these animals may be mosaics without the potential for germline transmission of the observed DNA modification. Furthermore, tissue extraction from B. mori larvae is an invasive procedure that can result in mortality. Therefore, we rear all G0 larvae without interference and begin molecular screening on G1 egg batches.

Initial Identification of Mutations
The initial G1 embryo screening for mutations ( Figure 2) involves a heteroduplex assay, which allows for the identification of the egg batch/es carrying promising mutations A workflow for the initial identification of mutations. (a) G 0 injected eggs are reared following microinjection and crossed with wildtype moths to generate (b) G 1 egg batches. (c) DNA extraction and PCR are performed on eggs from G 1 egg batches followed by (d) temperature denaturing and heteroduplex formation. (e) A PAGE-based heteroduplex assay is performed to identify which egg batches contain mutations. In this scheme G 0 _1 and G 0 _3 moths carry mutations in germline cells and give rise to mutant G 1 progeny. G 0 _2 moth carries somatic mutations which are not transmitted to the G 1 progeny. G 0 _4 lacks any mutation.
Molecular screening for the establishment of a stable homozygous gene knockout line involves three primary steps; (1) initial mutant screening with heteroduplex assay, (2) mutation sequence determination with TA cloning and Sanger sequencing, and (3) specific mutation screening by ARMS-PCR. Some authors begin initial screening on G 0 injected larvae; however, these animals may be mosaics without the potential for germline transmission of the observed DNA modification. Furthermore, tissue extraction from B. mori larvae is an invasive procedure that can result in mortality. Therefore, we rear all G 0 larvae without interference and begin molecular screening on G 1 egg batches.

Initial Identification of Mutations
The initial G 1 embryo screening for mutations ( Figure 2) involves a heteroduplex assay, which allows for the identification of the egg batch/es carrying promising mutations and is based on PCR amplification and PAGE. There is an option here on the approach that can be taken and should be determined by the number of putative mutated G 1 egg batches that are being analyzed (see Notes 1 and 2, Section 3.1.1).

Mutation Sequence Determination
The molecular characterization of the mutations is performed using TA cloning and Sanger sequencing on G 1 embryo DNA ( Figure 3). This step allows for the determination of the actual nature of the mutation/s and the designing of suitable primers required for the subsequent ARMS-PCR screening step (Sections 2.3 and 3.2). Molecular characterization requires PCR amplification, cloning into a TA cloning vector followed by Sanger sequencing and basic computational analyses, then designing appropriate primers to amplify a 400 to 1000 bp region around the genomic target. The primer couples employed to verify the gRNAs specificity in vitro can be used at this stage. Cloning requires both a cloning vector system and competent bacterial cells. We have used the StrataClone PCR Cloning Kit, which ligates PCR amplicons into a TA cloning vector that are transformed with kit specific competent bacteria. For sequencing, many companies provide plasmid primers with optimized running parameters. If this option is not available, it is possible to employ the PCR primers used for cloning. Subsequently, a sequence alignment must be performed to identify each specific mutation and determine whether a stop codon will be produced in the protein sequence. These computational analyses can be completed with a large number of software packages. We describe the process (Supplementary Methods A) with the use of three freely available software packages; CLC Sequence Viewer (V.8, Qiagen), MEGA 7 and Serial Cloner. and is based on PCR amplification and PAGE. There is an option here on the approach that can be taken and should be determined by the number of putative mutated G1 egg batches that are being analyzed (see Notes 1 and 2, Section 3.1.1).

Mutation Sequence Determination
The molecular characterization of the mutations is performed using TA cloning and Sanger sequencing on G1 embryo DNA ( Figure 3). This step allows for the determination of the actual nature of the mutation/s and the designing of suitable primers required for the subsequent ARMS-PCR screening step (Sections 2.3 and 3.2). Molecular characterization requires PCR amplification, cloning into a TA cloning vector followed by Sanger sequencing and basic computational analyses, then designing appropriate primers to amplify a 400 to 1000 bp region around the genomic target. The primer couples employed to verify the gRNAs specificity in vitro can be used at this stage. Cloning requires both a cloning vector system and competent bacterial cells. We have used the StrataClone PCR Cloning Kit, which ligates PCR amplicons into a TA cloning vector that are transformed with kit specific competent bacteria. For sequencing, many companies provide plasmid primers with optimized running parameters. If this option is not available, it is possible to employ the PCR primers used for cloning. Subsequently, a sequence alignment must be performed to identify each specific mutation and determine whether a stop codon will be produced in the protein sequence. These computational analyses can be completed with a large number of software packages. We describe the process (Supplementary Methods A) with the use of three freely available software packages; CLC Sequence Viewer (V.8, Qiagen), MEGA 7 and Serial Cloner.

Hemolymph Sampling and Screening with ARMS-PCR
Screening for heterozygous, homozygous, and hemizygous mutants is comprised of (i) hemolymph sampling from larvae, (ii) DNA extraction, and (iii) screening by ARMS-PCR, which allows for the identification of the mutation-bearing organisms ( Figure 4). These are then selected for the generation of the stable mutant line.

Hemolymph Sampling and Screening with ARMS-PCR
Screening for heterozygous, homozygous, and hemizygous mutants is comprised of (i) hemolymph sampling from larvae, (ii) DNA extraction, and (iii) screening by ARMS-PCR, which allows for the identification of the mutation-bearing organisms ( Figure 4). These are then selected for the generation of the stable mutant line.  Hemolymph sampling is a critical step since it may have deleterious effects on larval development, increasing the mortality rate. However, in trialing non-invasive DNA extraction methods using frass (feces) and exuviae (cast-off outer skins of the larvae at the molting stage) we achieved poor and inconsistent results. Here we describe our optimized protocol to extract B. mori hemolymph minimizing mortality. Regarding the DNA extraction procedure, we found that the PureYield Plasmid Miniprep System (Promega, Dane County, WI, USA) provides the highest quality and quantity of DNA (600 ng when eluted in 30 μL of H2O) from 20 μL of hemolymph. We also achieved good results using the Wizard ® Genomic DNA Purification Kit (Promega, Dane County, WI, USA) and the GeneJet Genomic DNA purification kit (ThermoFisher, Waltham, MA, USA), while a 10% Chelex buffer did not produce reproducible outcomes. Following sampling, G1 larvae must be reared individually; we find Petri dishes ideal housing as the larvae can be observed without disturbance and are easy to stack, secure, and transport. The ARMS-PCR method screens for known mutations by PCR and electrophoresis [16]. ARMS-PCR requires two PCR reactions for each sample, one containing amplification primers for the wildtype sequence and the second containing primers specifically designed to amplify the mutant haplotype (e.g., a modified forward primer bearing the mutated sequence at its 3′ region). For each reaction, it is suggested to add an internal PCR control, co-amplifying any genomic region easily distinguishable from the ARMS-PCR amplicons on an agarose gel. We amplify a 500 bp sequence of the gene cycle (cyc) as an internal control. ARMS-PCR is repeated with G2 animals, selecting hemi-or homozygous mutant females and males, which will establish the stable mutant line (from G3 onward) ( Figure 5). Hemolymph sampling is a critical step since it may have deleterious effects on larval development, increasing the mortality rate. However, in trialing non-invasive DNA extraction methods using frass (feces) and exuviae (cast-off outer skins of the larvae at the molting stage) we achieved poor and inconsistent results. Here we describe our optimized protocol to extract B. mori hemolymph minimizing mortality. Regarding the DNA extraction procedure, we found that the PureYield Plasmid Miniprep System (Promega, Dane County, WI, USA) provides the highest quality and quantity of DNA (600 ng when eluted in 30 µL of H 2 O) from 20 µL of hemolymph. We also achieved good results using the Wizard ® Genomic DNA Purification Kit (Promega, Dane County, WI, USA) and the GeneJet Genomic DNA purification kit (ThermoFisher, Waltham, MA, USA), while a 10% Chelex buffer did not produce reproducible outcomes. Following sampling, G 1 larvae must be reared individually; we find Petri dishes ideal housing as the larvae can be observed without disturbance and are easy to stack, secure, and transport. The ARMS-PCR method screens for known mutations by PCR and electrophoresis [16]. ARMS-PCR requires two PCR reactions for each sample, one containing amplification primers for the wildtype sequence and the second containing primers specifically designed to amplify the mutant haplotype (e.g., a modified forward primer bearing the mutated sequence at its 3 region). For each reaction, it is suggested to add an internal PCR control, co-amplifying any genomic region easily distinguishable from the ARMS-PCR amplicons on an agarose gel. We amplify a 500 bp sequence of the gene cycle (cyc) as an internal control. ARMS-PCR is repeated with G 2 animals, selecting hemi-or homozygous mutant females and males, which will establish the stable mutant line (from G 3 onward) ( Figure 5). wildtype sequence and the second containing primers specifically designed to amplify the mutant haplotype (e.g., a modified forward primer bearing the mutated sequence at its 3′ region). For each reaction, it is suggested to add an internal PCR control, co-amplifying any genomic region easily distinguishable from the ARMS-PCR amplicons on an agarose gel. We amplify a 500 bp sequence of the gene cycle (cyc) as an internal control. ARMS-PCR is repeated with G2 animals, selecting hemi-or homozygous mutant females and males, which will establish the stable mutant line (from G3 onward) ( Figure 5).  1.
Following CRISPR/Cas9 microinjection, rear the G 0 individuals until adult stage as in [23,24] and outbreed singly with wildtype moths from the same genetic background. After egg-laying, label each egg batch with a unique code.

2.
Allow the G 1 eggs to develop for three days, collect five eggs per egg batches, place each egg into a microtube. Label microtubes with the unique egg batch code, and replicate number (see Note 1).

3.
Crush the eggs with a pipette tip and perform DNA extraction and purification using the chosen DNA purification kit following the manufacturer's instructions. Quantify DNA with a NanoDrop and prepare 20 ng/µL working solutions for each sample.

4.
Methods Protoc. 2021, 4, x FOR PEER REVIEW 10 of 14 4. On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles. 5. Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish. 6. Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity ( Figure 6b), a drop of hemolymph will pool at the wound ( Figure 6c). 7.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette ( Figure  6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 8. Place the larva into the appropriately labeled Petri dish. Continue sampling remaining larvae. Allow the sampled larvae heal for a minimum of 20 min. After healing, feed larvae in the Petri dishes. 9.
PAUSE STEP Hemolymph samples can be frozen at −20 °C until use. 10. To rear sampled G1 larvae: everyday, remove the Petri dish lid and place upside down. Using forceps, transfer the larva to the lid, empty the frass and dry food from the dish, if necessary wipe the dish with ethanol soaked lab tissue before transferring the larva back with fresh food. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, add a folded square of lab tissue to facilitate cocoon spinning. After spinning begins do not interfere with the larvae for 4 days, then the pupae can be cut from the cocoons and sexed (Figure 6e). 3.3.2. DNA Extraction from Hemolymph and Screening with ARMS-PCR CRITICAL STEP Also perform DNA extraction on wildtype tissue from the same genetic background. This DNA will be used to obtain amplicons required for the mixed heteroduplex assay.

5.
Methods Protoc. 2021, 4, x FOR PEER REVIEW 10 of 14 4. On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles. 5. Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish. 6. Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity (Figure 6b), a drop of hemolymph will pool at the wound (Figure 6c). 7.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette ( Figure  6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 8. Place the larva into the appropriately labeled Petri dish. Continue sampling remaining larvae. Allow the sampled larvae heal for a minimum of 20 min. After healing, feed larvae in the Petri dishes. 9.
PAUSE STEP Hemolymph samples can be frozen at −20 °C until use. 10. To rear sampled G1 larvae: everyday, remove the Petri dish lid and place upside down. Using forceps, transfer the larva to the lid, empty the frass and dry food from the dish, if necessary wipe the dish with ethanol soaked lab tissue before transferring the larva back with fresh food. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, add a folded square of lab tissue to facilitate cocoon spinning. After spinning begins do not interfere with the larvae for 4 days, then the pupae can be cut from the cocoons and sexed (Figure 6e). (e) morphological characteristics used to sex pupae, females bear a vertical line on the penultimate (8th) segment, males bear a whole penultimate segment and have a CRITICAL STEP Use high fidelity polymerase when performing initial molecular screening. Calculate a sufficient PCR master mix volume for all your samples in 20 µL PCR reactions, include negative and positive controls. Calculate the required volume of wildtype amplicon (for heteroduplex formation, a minimum of 10 µL of wildtype amplicon per sample: e.g., 10 µL × 12 samples = 120 µL).

6.
Prepare a PCR master mix (see primer sequences in Figure 1 and Table 1), for 480 µL add reagents as in Table 2 4. On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles. 5. Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish. 6. Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity (Figure 6b), a drop of hemolymph will pool at the wound (Figure 6c). 7.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette ( Figure  6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 8. Place the larva into the appropriately labeled Petri dish. Continue sampling remaining larvae. Allow the sampled larvae heal for a minimum of 20 min. After healing, feed larvae in the Petri dishes. 9.
PAUSE STEP Hemolymph samples can be frozen at −20 °C until use. 10. To rear sampled G1 larvae: everyday, remove the Petri dish lid and place upside down. Using forceps, transfer the larva to the lid, empty the frass and dry food from the dish, if necessary wipe the dish with ethanol soaked lab tissue before transferring the larva back with fresh food. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, add a folded square of lab tissue to facilitate cocoon spinning. After spinning begins do not interfere with the larvae for 4 days, then the pupae can be cut from the cocoons and sexed (Figure 6e). CRITICAL STEP High fidelity polymerase cannot be used for ARMS-PCR. Prepare PCR master mixes for both wildtype (WT) and mutant (M) ARMS-PCR reactions as in Table 4. 4. On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles. 5. Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish. 6. Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity (Figure 6b), a drop of hemolymph will pool at the wound (Figure 6c). 7.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette ( Figure  6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 8. Place the larva into the appropriately labeled Petri dish. Continue sampling remaining larvae. Allow the sampled larvae heal for a minimum of 20 min. After healing, feed larvae in the Petri dishes. 9.
PAUSE STEP Hemolymph samples can be frozen at −20 °C until use. 10. To rear sampled G1 larvae: everyday, remove the Petri dish lid and place upside down. Using forceps, transfer the larva to the lid, empty the frass and dry food from the dish, if necessary wipe the dish with ethanol soaked lab tissue before transferring the larva back with fresh food. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, add a folded square of lab tissue to facilitate cocoon spinning. After spinning begins do not interfere with the larvae for 4 days, then the pupae can be cut from the cocoons and sexed (Figure 6e). . Load an appropriate DNA ladder, negative PCR control, and a positive control (wildtype only PCR amplicon). Run the gel at 50 V for 1 h, then increase to 200 V for 2.5 h. After the run, remove the gel from the plates, wash it three times with distilled water, submerge the gel in 75-100 mL of distilled water and add 5 µL of ethidium bromide (10 mg/mL), gently shake for 15 min. Image the gel on a Gel Doc™ XR+.
Note 1: With four or more egg batches, it will be more efficient to first screen pooled G 1 egg batches, by combining five to ten eggs in one microtube, and following steps 3, 4, 6, 7 and Section 3.1.2. Then select the specific egg batches bearing DNA modifications and follow this protocol from step 3, using single eggs as samples.
Note 2: As an alternative to the heteroduplex assay direct sequencing of the PCR products and analysis by using specific software [e.g., Tracking of Indel by Decomposition (TIDE https://tide.nki.nl/ [25]) or ICE Analysis (https://ice.synthego.com/) [26]] is possible. These analyses provide computational readouts allowing the identification of egg batches carrying mutant G 1 individuals. Carefully evaluate which approach to take depending on the number of samples, as direct sequencing and computational analysis are less labor-intensive but more time and cost demanding compared to heteroduplex assay.

Mutant Sequence Determination
PCR, Cloning and Sanger sequencing. Time for Completion: PCR-3 h; TA cloning 1.
Perform PCR using standard Taq polymerase and the gDNA previously extracted from the known mutated G 1 egg samples (see point 2 in Section 3.1.1). Prepare 480 µL of DreamTaq master mix as in Table 3 (see primer sequences in Figure 1 and Table 1 4. On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles. 5. Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish. 6. Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity (Figure 6b), a drop of hemolymph will pool at the wound (Figure 6c). 7.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette ( Figure  6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 8. Place the larva into the appropriately labeled Petri dish. Continue sampling remaining larvae. Allow the sampled larvae heal for a minimum of 20 min. After healing, feed larvae in the Petri dishes. 9.
PAUSE STEP Hemolymph samples can be frozen at −20 °C until use. 10. To rear sampled G1 larvae: everyday, remove the Petri dish lid and place upside down. Using forceps, transfer the larva to the lid, empty the frass and dry food from the dish, if necessary wipe the dish with ethanol soaked lab tissue before transferring the larva back with fresh food. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, add a folded square of lab tissue to facilitate cocoon spinning. After spinning begins do not interfere with the larvae for 4 days, then the pupae can be cut from the cocoons and sexed (Figure 6e). CRITICAL STEP High fidelity polymerase cannot be used for ARMS-PCR. Prepare PCR master mixes for both wildtype (WT) and mutant (M) ARMS-PCR reactions as in Table 4. PAUSE STEP Ligated plasmids can be stored as a reaction mixture at −20 • C before transformation.

5.
Transform competent cells following the provided protocol and incubate ON at 37 • C. 6.
Select at least five colonies from each ON plate.  4. On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles. 5. Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish. 6. Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity (Figure 6b), a drop of hemolymph will pool at the wound (Figure 6c). 7.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette ( Figure  6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 8. Place the larva into the appropriately labeled Petri dish. Continue sampling remaining larvae. Allow the sampled larvae heal for a minimum of 20 min. After healing, feed larvae in the Petri dishes. 9.
PAUSE STEP Hemolymph samples can be frozen at −20 °C until use. 10. To rear sampled G1 larvae: everyday, remove the Petri dish lid and place upside down. Using forceps, transfer the larva to the lid, empty the frass and dry food from the dish, if necessary wipe the dish with ethanol soaked lab tissue before transferring the larva back with fresh food. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, add a folded square of lab tissue to facilitate cocoon spinning. After spinning begins do not interfere with the larvae for 4 days, then the pupae can be cut from the cocoons and sexed (Figure 6e).
CRITICAL STEP Test the ARMS-PCR primers using wildtype gDNA. Mutant (M)-ARMS-PCR primers must not generate an amplicon using wildtype gDNA. For each positive G1 batch, process at least 30-50 larvae.

3.
Sterilize the workspace with 70% ethanol. For each larva, label one 1.5 mL microtube and one sterile Petri dish with the same code. Repeat for each larva, in advance. Place the labeled microtubes on ice, on the workbench near the labeled Petri dishes.

4.
On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles.

5.
Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish.

6.
Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity (Figure 6b), a drop of hemolymph will pool at the wound (Figure 6c).

7.
4. On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles. 5. Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish. 6. Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity (Figure 6b), a drop of hemolymph will pool at the wound (Figure 6c). 7.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette ( Figure  6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 8. Place the larva into the appropriately labeled Petri dish. Continue sampling remaining larvae. Allow the sampled larvae heal for a minimum of 20 min. After healing, feed larvae in the Petri dishes. 9.
PAUSE STEP Hemolymph samples can be frozen at −20 °C until use. 10. To rear sampled G1 larvae: everyday, remove the Petri dish lid and place upside down. Using forceps, transfer the larva to the lid, empty the frass and dry food from the dish, if necessary wipe the dish with ethanol soaked lab tissue before transferring the larva back with fresh food. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, add a folded square of lab tissue to facilitate cocoon spinning. After spinning begins do not interfere with the larvae for 4 days, then the pupae can be cut from the cocoons and sexed (Figure 6e). CRITICAL STEP High fidelity polymerase cannot be used for ARMS-PCR. Prepare PCR master mixes for both wildtype (WT) and mutant (M) ARMS-PCR reactions as in Table 4.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette (Figure 6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 8.
Place the larva into the appropriately labeled Petri dish. Continue sampling remaining larvae. Allow the sampled larvae heal for a minimum of 20 min. After healing, feed larvae in the Petri dishes. 9.
4. On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles. 5. Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish. 6. Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity (Figure 6b), a drop of hemolymph will pool at the wound (Figure 6c). 7.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette ( Figure  6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 8. Place the larva into the appropriately labeled Petri dish. Continue sampling remaining larvae. Allow the sampled larvae heal for a minimum of 20 min. After healing, feed larvae in the Petri dishes. 9.
PAUSE STEP Hemolymph samples can be frozen at −20 °C until use. 10. To rear sampled G1 larvae: everyday, remove the Petri dish lid and place upside down. Using forceps, transfer the larva to the lid, empty the frass and dry food from the dish, if necessary wipe the dish with ethanol soaked lab tissue before transferring the larva back with fresh food. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, add a folded square of lab tissue to facilitate cocoon spinning. After spinning begins do not interfere with the larvae for 4 days, then the pupae can be cut from the cocoons and sexed (Figure 6e). CRITICAL STEP High fidelity polymerase cannot be used for ARMS-PCR. Prepare PCR master mixes for both wildtype (WT) and mutant (M) ARMS-PCR reactions as in Table 4. PAUSE STEP Hemolymph samples can be frozen at −20 • C until use. 10. To rear sampled G 1 larvae: everyday, remove the Petri dish lid and place upside down. Using forceps, transfer the larva to the lid, empty the frass and dry food from the dish, if necessary wipe the dish with ethanol soaked lab tissue before transferring the larva back with fresh food. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, add a folded square of lab tissue to facilitate cocoon spinning. After spinning begins do not interfere with the larvae for 4 days, then the pupae can be cut from the cocoons and sexed (Figure 6e).
4. On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles. 5. Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish. 6. Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity (Figure 6b), a drop of hemolymph will pool at the wound (Figure 6c). 7.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette ( Figure  6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 8. Place the larva into the appropriately labeled Petri dish. Continue sampling remaining larvae. Allow the sampled larvae heal for a minimum of 20 min. After healing, feed larvae in the Petri dishes. 9.
PAUSE STEP Hemolymph samples can be frozen at −20 °C until use. 10. To rear sampled G1 larvae: everyday, remove the Petri dish lid and place upside down. Using forceps, transfer the larva to the lid, empty the frass and dry food from the dish, if necessary wipe the dish with ethanol soaked lab tissue before transferring the larva back with fresh food. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, add a folded square of lab tissue to facilitate cocoon spinning. After spinning begins do not interfere with the larvae for 4 days, then the pupae can be cut from the cocoons and sexed (Figure 6e). CRITICAL STEP High fidelity polymerase cannot be used for ARMS-PCR. Prepare PCR master mixes for both wildtype (WT) and mutant (M) ARMS-PCR reactions as in Table 4. 4. On the workbench, prepare; a small beaker with 50 mL of 70% ethanol, lab tissue, a microtube rack, a P20 micropipette, sterile tips, hypodermic needles, and waste containers for general lab waste, and hypodermic needles.
5. Using forceps, select a larva and carefully submerge it into the 70% ethanol beaker for 3 s. Gently blot the larva dry on lab tissue and let air dry for a few seconds. While air drying, open a labeled microtube and take its paired Petri dish.
6. Pick up and gently fold the worm head-to-tail, exposing the dermis between the body folds ( Figure 6a). Hold the larva over the microtube and use a hypodermic needle to gently pierce the dermis without deeply entering the body cavity (Figure 6b), a drop of hemolymph will pool at the wound (Figure 6c).

7.
CRITICAL STEP Collect the hemolymph drop using a P20 micropipette ( Figure   6d) and place into the microtube. Do not squeeze the larvae or attempt to withdraw additional hemolymph from the wound with the micropipette. Store the hemolymph on ice. 11. After 6-7 days the larvae will prepare to spin their cocoon. Empty and clean the dish, CRITICAL STEP High fidelity polymerase cannot be used for ARMS-PCR. Prepare PCR master mixes for both wildtype (WT) and mutant (M) ARMS-PCR reactions as in Table 4. Rear G 2 larvae until the 5th instar and perform DNA extraction from hemolymph and ARMS-PCR as from Section 3.3.1.

7.
Select hemi-and homozygous mutant G 2 larvae and rear until adult stage. Mate them to generate a G 3 mutant stable line.

Initial Identification of Mutations
Successful PCR of the target region should show a clear single band in a 1.2% agarose gel (Figure 7a). Following heteroduplex formation by temperature ramp-down, the PCR products are analyzed by PAGE. G 1 eggs carrying a mutation can be identified from the interpretation of the heteroduplex assay results. In fact, wildtype samples and the positive control will show a single band, corresponding to a homoduplex fragment (Figure 7b). Samples carrying a mutation will appear with larger multiband, corresponding to heteroduplexes (Figure 7b). DNA mutant samples are then selected for subsequent cloning and Sanger sequencing.

Initial Identification of Mutations
Successful PCR of the target region should show a clear single band in a 1.2 % agarose gel (Figure 7a). Following heteroduplex formation by temperature ramp-down, the PCR products are analyzed by PAGE. G1 eggs carrying a mutation can be identified from the interpretation of the heteroduplex assay results. In fact, wildtype samples and the positive control will show a single band, corresponding to a homoduplex fragment (Figure 7b). Samples carrying a mutation will appear with larger multiband, corresponding to heteroduplexes (Figure 7b). DNA mutant samples are then selected for subsequent cloning and Sanger sequencing.

Mutant Sequence Determination
Following cloning, Sanger sequencing will return either wildtype or mutated sequences (Figure 8a). The identified mutation is then analyzed to observe whether a stop codon is generated in the protein-coding sequence (Figure 8b).

Mutant Sequence Determination
Following cloning, Sanger sequencing will return either wildtype or mutated sequences (Figure 8a). The identified mutation is then analyzed to observe whether a stop codon is generated in the protein-coding sequence (Figure 8b). As a note, PCR samples derived from G1 embryos could also be processed with direct Sanger sequencing. Homozygous wildtypes will generate clear single peaks throughout the chromatogram, while heterozygous mutant samples will show double peaks downstream of the PAM site (Figure 8c). It is important to mention that genes targeted on the Z sex chromosome will result in hemizygous females either positive or negative for the mutation. Samples from hemizygous females appear identical to homozygous samples (either wildtype or mutant).

Screening with ARMS-PCR
The mutant and wildtype sequences are used to design ARMS-PCR primers. We designed two forward primers, ARMS_WT_FOR and ARMS_M_FOR respectively able to amplify the wildtype or mutant sequences, when used with the same reverse per_REV primer ( Table 1). The expected banding patterns for successful ARMS-PCR are shown in Figure  9. All samples will show a band corresponding to the internal PCR control (cyc amplicon). A per wildtype sample will produce a per amplification band in the WT-ARMS reaction and no per amplification bands in the M-ARMS reaction. A heterozygous larva will produce per amplification bands in both reactions. In contrast, a homozygous mutant will produce no per amplification bands in the WT-ARMS reaction and a per amplification band in the M-ARMS reaction. As per is located on the Z sex chromosome, females will be hemizygous wildtype or mutants, only displaying a single band in either the wildtype or mutant ARMS-PCR, respectively. As a note, PCR samples derived from G 1 embryos could also be processed with direct Sanger sequencing. Homozygous wildtypes will generate clear single peaks throughout the chromatogram, while heterozygous mutant samples will show double peaks downstream of the PAM site ( Figure 8c). It is important to mention that genes targeted on the Z sex chromosome will result in hemizygous females either positive or negative for the mutation. Samples from hemizygous females appear identical to homozygous samples (either wildtype or mutant).

Screening with ARMS-PCR
The mutant and wildtype sequences are used to design ARMS-PCR primers. We designed two forward primers, ARMS_WT_FOR and ARMS_M_FOR respectively able to amplify the wildtype or mutant sequences, when used with the same reverse per_REV primer ( Table 1). The expected banding patterns for successful ARMS-PCR are shown in Figure 9. All samples will show a band corresponding to the internal PCR control (cyc amplicon). A per wildtype sample will produce a per amplification band in the WT-ARMS reaction and no per amplification bands in the M-ARMS reaction. A heterozygous larva will produce per amplification bands in both reactions. In contrast, a homozygous mutant will produce no per amplification bands in the WT-ARMS reaction and a per amplification band in the M-ARMS reaction. As per is located on the Z sex chromosome, females will be hemizygous wildtype or mutants, only displaying a single band in either the wildtype or mutant ARMS-PCR, respectively. Methods Protoc. 2021, 4, x FOR PEER REVIEW 13 of 14 To establish a stable per mutant line, only G1 females bearing a per amplicon in the M-ARMS-PCR reaction and G1 males bearing a per amplicon in both ARMS-PCR reactions should be selected, followed until adult stage, and crossed to generate G2 progenies. G2 5th instar larvae will be screened via hemolymph DNA extraction and ARMS-PCR, identifying the appropriate hemizygous and homozygous mutant individuals to -found the stable mutant line, from G3 onward.

Conclusions
CRISPR/Cas9 is an efficient and relatively simple gene-editing technology rapidly becoming the technique of choice for the generation of novel gene knock-out and knockin research organisms. A variety of screening methods are available to determine the nature of CRISPR/Cas9 induced mutations and mutant baring individuals, and screening workflows should be tailored to specific target species. Here, we have reported an efficient screening protocol to identify CRISPR/Cas9-induced indels in a Z-linked gene in the silkworm Bombyx mori and have applied to generate a stable mutant line. This workflow can be employed with standard molecular biology equipment and can be generally used to identify a wide range of CRISPR/Cas9-mediated mutations within genes localized on both autosomes and heteromorphic sex chromosomes. This method could also be extended to detect CRISPR/Cas9-mediated mutations in other model and non-model insects, including haploid insects. To establish a stable per mutant line, only G 1 females bearing a per amplicon in the M-ARMS-PCR reaction and G 1 males bearing a per amplicon in both ARMS-PCR reactions should be selected, followed until adult stage, and crossed to generate G 2 progenies. G 2 5th instar larvae will be screened via hemolymph DNA extraction and ARMS-PCR, identifying the appropriate hemizygous and homozygous mutant individuals to -found the stable mutant line, from G 3 onward.

Conclusions
CRISPR/Cas9 is an efficient and relatively simple gene-editing technology rapidly becoming the technique of choice for the generation of novel gene knock-out and knockin research organisms. A variety of screening methods are available to determine the nature of CRISPR/Cas9 induced mutations and mutant baring individuals, and screening workflows should be tailored to specific target species. Here, we have reported an efficient screening protocol to identify CRISPR/Cas9-induced indels in a Z-linked gene in the silkworm Bombyx mori and have applied to generate a stable mutant line. This workflow can be employed with standard molecular biology equipment and can be generally used to identify a wide range of CRISPR/Cas9-mediated mutations within genes localized on both autosomes and heteromorphic sex chromosomes. This method could also be extended to detect CRISPR/Cas9-mediated mutations in other model and non-model insects, including haploid insects.