Design and Experiments

This section describes the experimental design and protocols for

1. The sgRNA plasmid cloning process
2. The RBD plasmid cloning process
3. The RBD swapping workflow

sgRNA Expression Plasmids

Four plasmids were tested for sgRNA cloning (as part of an iterative troubleshooting cycle, as described in the engineering page). Each plasmid contains an antibiotic resistance cassette and supports CRISPR-mediated editing through either Cas protein expression or sgRNA expression. pHERD30T-cas9 (abbreviated tcas9; Addgene 194476) and pCas3cRh (Addgene 133773) plasmids express gentamicin resistance (GmR), pCas9 (Addgene 42876) expresses chloramphenicol resistance (CmR), and pCRISPR (Addgene 113252) expresses ampicillin resistance (AmpR). Notably, pCas9 and pCas3cRh encode Cas nucleases (Cas9 and Cas3, respectively), while pCRISPR is dedicated to sgRNA expression and requires Cas9 supplied from a companion plasmid (typically pCas9). This is shown in Fig. 1 below. Four plasmids were tested for sgRNA cloning (as part of an iterative troubleshooting cycle, as described in the engineering page). Each plasmid contains an antibiotic resistance cassette and supports CRISPR-mediated editing through either Cas protein expression or sgRNA expression. pHERD30T-cas9 (abbreviated tcas9; Addgene 194476) and pCas3cRh (Addgene 133773) plasmids express gentamicin resistance (GmR), pCas9 (Addgene 42876) expresses chloramphenicol resistance (CmR), and pCRISPR (Addgene 113252) expresses ampicillin resistance (AmpR). Notably, pCas9 and pCas3cRh encode Cas nucleases (Cas9 and Cas3, respectively), while pCRISPR is dedicated to sgRNA expression and requires Cas9 supplied from a companion plasmid (typically pCas9). This is shown in Fig. 1 below.

In general, the cloning processes for inserting the sgRNA sequences into these plasmids involved annealing forward (F) and reverse (​R) DNA oligos together (both of which code for the sgRNA). Both of these have complimentary overhangs to restriction sites within the backbone, allowing them to be easily ligated into a given expression plasmid. For certain cloning workflows (tCas9), the F and R oligos were first constructed using SOE-PCR and then cloned. For others (pCas3Rh, pCas9, pCRISPR), the F and R oligos were ordered as full constructs and then cloned.

tcas9 sgRNA cloning protocol

The Forward (F) and Reverse (​R) sgRNA oligos were added to a tube in equal molar amounts of DNA to total approximately 100 ng. Milli-Q water was added to achieve a total volume of 5 μL. 20 μL of Master Mix A given by the recipe from Table 1 was added and the first 10 steps of SOE PCR were run as seen in the SOE PCR program listed below. 25 μL of Master Mix B given by the recipe from Table 1 was added. This mix contains the primers that are needed for the reaction to take place. Finally, the SOE PCR program was run to completion and the assembled sgRNA was digested with Nhel-HF and PCR cleaned.

Reagent	Master Mix A	Master Mix B
Q5 2x Master Mix	12.5 μL	12.5 μL
10 μM External Forward Primer	0 μL	1.25 μL
10 μM External Reverse Primer	0 μL	1.25 μL
Sterile Milli-Q Water	7.5 μL	10μL

Table 1. The Master Mix recipe for SOE PCR.

SOE PCR Program (a typical program for a desired product less than 2 kb in size):

1. 94 °C - 5 min
2. 94 °C - 30 sec
3. 60 °C - 1.5 min
4. 72 °C - 2.5 min
5. Repeat steps 2-4 10X
6. Hold at 10 °C - 10 min (window of time to add mix B)
7. 94 °C - 30 sec
8. 58 °C - 30 sec
9. 72 °C - 15 sec
10. Repeat steps 7-9 35X
11. Hold at 72 °C - 10 min
12. Hold at 10 °C indefinitely

IC-CRISPR sgRNA assembly

IC-CRISPR refers to the cloning process involving pCas3Rh. Upon preparing the sgRNA for ligation, the forward and reverse oligos were annealed and phosphorylated, as the sgRNA was ordered single-stranded for cost efficiency. The oligo annealing was done in a 1.5 mL microfuge tube with the reagents shown in Table 2 below (for a reaction totaling 50 μL).

Reagent	Volume
Oligo I	1 μL
Oligo II	1 μL
10X T4 Ligase Buffer (NEB)	5 μL
T4 PNK (NEB) T4 Polynucleotide Kinase	1 μL
ddH20	42 μL

Table 2: Reagents and amounts used for annealing IC-CRISPR oligos.

The annealing reaction mix was incubated for 1 hour at 37 °C. Then, 2.5 μL of 1M NaCl was added to phosphorylate the annealed oligos.

Upon preparing the pCas3cRh plasmid backbones to receive the insert, the pCas3cRh plasmids were liquid cultured with GmR and miniprepped to isolate the plasmid. Miniprepping was done following the Monarch® Spin Plasmid Miniprep Kit protocols. Then, the plasmid was digested with BsaI-HF, a Type IIS restriction enzyme for Golden Gate Assembly (GGA). BsaI cuts the plasmid 1 bp outside from the BsaI restriction site, leaving a 4 bp overhang. The digestion protocol used the 50 μL reaction mixture in Table 3.

Reagent	Volume
pcas3cRh	X μL (0.5 μg to 2 μg of plasmid)
BsaI	1 μL (up to 2 μL)
10X Cutsmart Buffer	5 μL
ddH2O	Y μL (to bring up to 50 μL)

Table 3: Reagents and amounts used for digesting the pCas3cRh plasmid.

The reaction mixture was incubated at 37 °C for 1 hour, after which the digestion was PCR cleaned using the Monarch® PCR & DNA Cleanup Kit and eluted with 30 μL of ddH₂O. After preparing the sgRNA and plasmid backbone, ligation was performed to insert the sgRNA into the plasmid backbone using the 10 μL ligation mixture in Table 4:

Reagent	Volume
BsaI digested pCas3cRh	X μL (~50 ng)
Phosphorylated and annealed oligos	0.5 μL
T4 DNA buffer	1 μL
T4 DNA ligase	0.5 μL
ddH2O	Y μL (to bring reaction to 10 μL)

Table 4: Reagents and amounts used for the ligation reaction.

Then, the ligation reaction mixture was left to incubate at room temperature for 2 hours or at 16 °C overnight.

After the ligation was performed, the recombinant plasmid was transformed into DH5α with GmR, liquid cultured, then miniprepped to isolate the cloned plasmid before sending it for Sanger sequencing to verify if the assembly was successful.

pCas9 and pCRISPR sgRNA assembly protocol

pCas9 and pCRISPR both contain BsaI sites for sgRNA cloning similar to pCas3cRh. Therefore, the cloning strategies are similar as well. The forward and reverse oligos were annealed and phosphorylated. The oligo annealing was done in a 1.5 mL microfuge tube with the reagents in Table 5 for a reaction totalling 50 μL.

Reagent	Volume
Oligo I	1 μL
Oligo II	1 μL
10X T4 Ligase Buffer (NEB)	5 μL
T4 PNK (NEB) T4 Polynucleotide Kinase	1 μL
ddH20	42 μL

Table 5: Reagents and amounts used for annealing IC-CRISPR oligos.

The annealing reaction mix was incubated for 1 hour at 37 °C. Then, 2.5 μL of 1M NaCl was added to phosphorylate the annealed oligos.

To prepare pCas9 and pCRISPR for ligation, the plasmid was digested with BsaI-HF, a Type IIS restriction enzyme for Golden Gate Assembly (GGA). BsaI cuts the plasmid 1 bp outside from the BsaI restriction site, leaving a 4 bp overhang. The digestion protocol used the 50 μL reaction mixture in Table 6.

Reagent	Volume
pcas9/pCRISPR	X μL (0.5 μg to 2 μg of plasmid)
BsaI	1 μL (up to 2 μL)
10X Cutsmart Buffer	5 μL
ddH2O	Y μL (to bring up to 50 μL)

Table 6: Reagents and amounts used for digesting the pCas3cRh plasmid.

The reaction mixture was incubated at 37 °C for 1 hour, after which the digestion was PCR cleaned using the Monarch® PCR & DNA Cleanup Kit and eluted with 30 μL of ddH2O.

After preparing the sgRNA and plasmid backbone, ligation was performed to insert the sgRNA into the plasmid backbone using the 10 μL ligation mixture in Table 4:

Reagent	Volume
BsaI digested pCas9/pCRISPR	X μL (~50 ng)
Phosphorylated and annealed oligos	0.5 μL
T4 DNA buffer	1 μL
T4 DNA ligase	0.5 μL
ddH2O	Y μL (to bring reaction to 10 μL)

Table 7: Reagents and amounts used for the pcas9/pCRISPR ligation reaction.

Then, the ligation reaction mixture was left to incubate at room temperature for 2 hours or at 16 °C overnight.

After the ligation was performed, the cloned plasmid was transformed into DH5α with GmR, liquid cultured, then miniprepped to isolate the plasmid before sending it for Sanger sequencing to verify if the assembly was successful.

RBD Plasmid Assembly

For delivery of the RBD sequences (with homology arms to the prophage) into E. coli K12, RBDs synthesized from dry lab were cloned into a plasmid called pETDuet-1 (Addgene 71146). We first transformed our pETDuet-1 backbone (~5420 bp) into competent E. coli DH5α cells using ampicillin plates and incubated them overnight at 37 °C. We then digested the miniprepped samples of pETDuet-1 using NcoI and HindIII to linearize our backbone. We then incubated them at 37 °C for at least 3 hours.

Reagent	Volume
Miniprepped pETDuet-1	X μL (~1000 ng)
HindIII	1 μL
NcoI	1 μL
Cutsmart buffer	5 μL
ddH2O	Y μL (to bring reaction to 50 μL)

Table 8: Reagents and amounts used for the digestion of pETDuet-1.

The resulting fragment from the cut is 74 bp. NcoI and HindIII are both type II restriction enzymes that leave sticky ends and 4-nucleotide overhangs, allowing us to later integrate our generated RBD inserts from dry lab to create our plasmid using Gibson Assembly. These inserts are around 200 bp and contain the new RBD sequence, flanked by homology arms (HAs) to the WT prophage, which are further flanked by overhangs that match our pETDuet-1 backbone (for simplicity, these overhangs can also be thought of as “plasmid HAs”).

After digestion, we CIP-treated our digested samples using Quick CIP to remove the 5’-phosphate groups from each end of the linearized backbone in order to prevent self-ligation from taking place. We then purified our digested samples to isolate our linearized pETDuet-1 backbones. After, we used Gibson Assembly to integrate our generated RBD inserts into our purified linearized backbones. This entire process is shown in Fig. 2 below, and the Gibson Assembly recipe is shown in Table 9 below.

Reagent	Volume
Digested pETDuet-1	X μL (100ng)
RBD insert	Y μL (3x molar equivalent of pETDuet-1
NEBuilder HiFi DNA Assembly Master Mix	10 μL
Nuclease-free H2O	Z μL (to bring reaction to 20 μL

Table 9: Reagents and amounts used for Gibson assembly

We then took our assembled plasmid with the new RBD inserts and transformed them into DH5α, and let the plates incubate overnight at 37 °C. Using the transformed cells, we then created liquid cultures and minipreps and ran an agarose gel to check if our assembly worked. For a positive result, we would expect to see a single band around the 6 kb mark using a 1 kb+ DNA ladder. Following confirmation of successful assemblies, we then made glycerol stocks of successful transformations and saved the successful minipreps for cotransformation and rebooting.

RBD Swapping

Cotransformation

Once all necessary plasmids are assembled (pETDuet-1 and appropriate sgRNA plasmids), we co-transform both into lysogenic E. coli K12 for RBD swapping to occur. Following confirmation of the swap with colony PCR, we then induce resulting cultures with mitomycin C to “reboot” the new phages.

To begin, the necessary plasmids are co-transformed into lysogenic E. coli K12 then plated and selected via selectable markers (i.e. GmR, AmpR, CmR, or a combination of multiple depending on the plasmids used). Transformed colonies are picked and cultured overnight at 37 °C. We then add 5 mL of culture to 50 mL of LB media under selection – throughout this entire process, RBD swapping occurs. Following this, we add in 5 μL of mitomycin C once the culture’s OD600 reaches approximately 0.4. Mitomycin C is an antibiotic which shocks the cells to activate transcription of the integrated phage genome. Cells are grown at 37 °C until the culture turns clear – indicating that the phages are active and have lysed the cells – after which we add chloroform. We shake the sample at 37 °C for 15 minutes, then centrifuge and collect the supernatant for titration and phage spotting.

Looking deeper into the swapping process, the wild type RBD is replaced with our generated RBD. Recombination occurs both naturally and Cas-assisted, with the sgRNA plasmids acting as counterselection. Specifically, if natural recombination doesn’t occur, the Cas protein creates double stranded cuts in the phage genome, selecting against K12 cells with WT prophage RBDs. This also induces a mechanism similar to homology-directed repair, in which the generated RBD is utilized to repair the cut, thus enhancing recombination efficiency and ensuring the swap is more likely to be successful.

To validate whether an RBP swap has successfully occurred, we created checking primers to run colony PCR on our co-transformed cells. Primers are designed to target both the RBD sequence and DNA outside of the RBD + HA region; this way, the bands for the amplified region on a gel is indicative of a successful swap, and can be discriminated from off-target binding of the checking primers to RBDs integrated in the pETDuet-1 plasmid. We also run colony PCR with primers designed to target the wild type RBD as a control. For this specific case, no band will appear on this gel for successful swaps.

Fig. 3 below showcases the mechanism behind this process.

Phage Spotting

Phage spotting is the final stage in the experimental workflow. It is done by pipetting small droplets (1.5 μL) of the phage solution over a lawn of bacteria at 10-fold increasing dilutions. If the bacteriophage is able to lyse the bacteria it is being tested against, a large plaque or various small plaques will form on the lawn, reflecting both the efficiency at which the phage lyses its target, and the concentration at which it is being tested (note: plaques are zones of clearance of bacteria on the agar plate). By obtaining a countable number of plaques, it is possible to quantify phage titres in plaque-forming units per milliliter (PFU/mL).

The goal of the phage spotting assay is to evaluate how host specificity changes when wild-type phage RBDs are replaced with dry lab-generated variants. We use two glycan-binding phages (Mu and P2) and two protein-binding phages (Lambda and HK97), alongside a panel of E. coli strains: K12, BL21, ΔOmpC, and ΔLamB. Wild-type Mu and P2 recognize the K12 LPS core, while Lambda and HK97 recognize the LamB receptor on K12. BL21 instead has a truncated R1-core LPS, preventing infection by Mu and P2, while ΔLamB lacks the LamB receptor needed for Lambda and HK97 infection. Conversely, ΔOmpC lacks OmpC, which is not normally used by wild-type protein-binding phages but is the intended target for recombinant Lambda and HK97. Thus, K12 and ΔOmpC serve as positive controls for wild-type glycan- and protein-binding phages respectively, while BL21 and ΔLamB serve as negative controls. Recombinant phages should switch specificity: Mu and P2 to BL21’s truncated R1 LPS, and Lambda and HK97 to the OmpC receptor in K12 (see Fig. 4 below).

Assembly Sequencing

Successfully transformed bacteria may have one of two plasmids: The original vector with no sgRNA insert, or assembled plasmid with sgRNA insert. To verify successful cloning of sgRNA inserts into their vectors, we employed Sanger sequencing at the insert site in the plasmid. Primers complementary to sites surrounding the insert sites were added, and the insert was amplified by a mixture of dNTPs and fluorescently labeled terminating ddNTPs, resulting in a mixture of PCR products with varying lengths. Capillary electrophoresis with fluorescence reading is used to determine the sequence of the amplified insert^[1]. Resulting DNA sequences acquired from fluorescence readings are aligned against expected vector assemblies and successful sgRNA assembly is verified by presence of a 34bp sequence (IC-CRISPR) or 40bp sequence (tCas9) at the site of insert analogous to the transformed sgRNA.

Lab Notes

BioZone and BME Lab Notes

MaRS Lab Notes