Funding for this work was provided by the Fondation Bettencourt Schueller in support of the Paris Bettencourt iGEM team. We thank the INSERM U1001 research unit and Chantal Lotton for technical assistance and advice. Phagemid Litmus28i_J23115-B0032-GFP was provided by Monica Ortiz and Drew Endy of Stanford.

1. Design and Construction of Phagemid Vectors Bearing sRNA Silencing Cassettes
<ol>
	<li>De Novo Design of sRNA Silencing Cassettes8
	<ol>
		<li>Identify the complete sequence of the mRNA to be silenced using a DNA sequence database. To generate the TARGET sequence, note the first 24 bp of coding sequence, from position +1 to +24 beginning with the start codon (e.g., ATG). 
		Note: Silencing is less efficient when other sites or segments of the mRNA are targeted8.</li>
		<li>Take the reverse complement of the TARGET sequence to produce the GUIDE sequence for the sRNA cassette. See Table 1 for examples of TARGET and GUIDE sequences for chloramphenicol acetyltransferase (CAT).</li>
		<li>To design the 292 bp complete sRNA expression cassette, arrange the pR promoter, GUIDE sequence, Hfq protein binding domain and T1/TE transcriptional terminator sequences in series (Table 2).</li>
		<li>Add additional cloning sites of choice to facilitate cloning of the sRNA cassette into the target vector.</li>
		<li>Obtain the complete sRNA cassette through commercial gene synthesis or a similar method and clone it into any phagemid vector with a functional f1 replication origin11. See Supporting Information for the complete sequence of the final phagemid vector.</li>
	</ol>
	</li>
	<li>Altering the Sequence Targeted by an Existing sRNA Expression Cassette using PCR-based Site Directed Mutagenesis12
	<ol>
		<li>Identify the 24 bp GUIDE sequence in an existing sRNA expression cassette. Note: The annotated pAB.001 plasmid, used in this work, is available as a supplemental sequence file.</li>
		<li>Design forward and reverse primers with short regions of homology to the existing sRNA cassette flanking the new 24 bp GUIDE sequence. Obtain the primers through commercial oligonucleotide synthesis. 
		Note: Primer design for site-directed mutagenesis is depicted in Figure 2. Exact primer template sequences are provided in Table 3.</li>
		<li>Prepare a 5 ml culture of E. coli carrying the template sRNA expression phagemid. Grow the cells overnight at 37 &#176;C with shaking, in LB media with appropriate antibiotics.</li>
		<li>Extract and purify the template sRNA expression phagemid from the 5 ml bacterial culture using a DNA miniprep kit or similar method12.</li>
		<li>Prepare two PCR reactions using the template sRNA expression phagemid and high fidelity polymerase, one with the forward and one with the reverse primer (Table 4). Use PCR conditions as recommended by the polymerase supplier (Table 5). Increase the template concentration to 10-50x higher than a standard reaction to account for the fact that the single primer reaction does not produce exponential amplification.</li>
		<li>Combine the two above PCR reactions in a microcentrifuge tube. Anneal the products by heating to 98 &#176;C in a boiling water bath. Immediately after placing the microcentrifuge tube in the water bath, remove the heat source and allow the bath to slowly return to room temperature over 1-2 hr.</li>
		<li>To eliminate the unmutated template sRNA expression phagemid, add 1 &#181;l. DpnI restriction enzyme to the mixture and incubate at 37 &#176;C for 1 hr, or the time recommended by the manufacturer for complete digestion. 
		Note: DpnI digests only methylated target sites, which are present on host-replicated phagemids but not PCR products.</li>
		<li>Transform purchased or prepared13 chemically competent E. coli with 1-5 &#181;l of the annealed PCR product. Isolate single colonies of the transformed strain by selective plating on LB agar plates containing appropriate antibiotics.</li>
		<li>To verify the incorporation of the correct GUIDE sequence, screen the resulting colonies by colony PCR. Using a 200 &#181;l pipette tip, collect a small quantity of cells from a single transformed colony. Mark and preserve the original colony for downstream use after verification.</li>
		<li>Add the collected cells to 50 &#181;l of nuclease-free water in a microcentrifuge tube. Mix by pipetting up and down.</li>
		<li>Using a bench-top thermocycler or a boiling water bath, lyse the cells by heating to 95 &#176;C for 2 min.</li>
		<li>PCR-amplify the phagemid region using 1 &#181;l of heat-lysed cells as a DNA template. PCR conditions and thermocycler protocols are provided in Tables 6 and 7. See the supplemental pAB.001 sequence file for verification primer sequences.</li>
		<li>Sequence the PCR product to verify incorporation of the correct GUIDE sequence.</li>
		<li>Inoculate a 5 ml culture of the E. coli clone carrying the sequence-verified sRNA expression phagemid. Grow the cells overnight at 37 &#176;C with shaking, in selective LB media.</li>
		<li>Prepare a glycerol stock of the sequence-verified clone. Add 750 &#181;l of the overnight culture to 250 &#181;l of 60% glycerol in a screw-cap cryo tube.</li>
		<li>Store the glycerol stock at -80 &#176;C indefinitely. The remainder of the overnight culture may be used as a source for the sRNA expression phagemid in step 2.</li>
	</ol>
	</li>
</ol>
2. Production and Harvest of M13-packaged Phagemid Stocks
<ol>
	<li>Prepare a 5 ml culture of E. coli carrying the sRNA expression phagemid. Grow the cells overnight at 37 &#176;C with shaking, in LB media with appropriate antibiotics. Note: The sRNA expression phagemid may be obtained through de novo cloning as described in step 1.1.5, or modified from an existing phagemid and harvested in step 1.2.16.</li>
	<li>Similarly prepare a 5 ml culture of E. coli carrying the M13KO7 helper plasmid. Grow the cells overnight at 37 &#176;C with shaking, in selective LB media.</li>
	<li>Extract and purify the sRNA expression phagemid and helper plasmid using a DNA extraction kit or similar method12.</li>
	<li>Cotransform purchased or prepared13 chemically competent E. coli with 1 &#181;l each of sRNA expression phagemid and helper plasmid. Select for cotransformants by plating on LB agar with selective antibiotics for both constructs.</li>
	<li>Prepare a 10 ml culture from a single colony of the cotransformed strain in LB with selective antibiotics. Incubate at 37 &#176;C with shaking for 8-12 hr or overnight.</li>
	<li>Centrifuge the culture at 3,300 x g for 10 min. Collect the supernatant and filter through a 0.2 &#181;m filter. Caution: In case of media spill, clean the area with dilute bleach (0.5%) to destroy infective phage particles.</li>
	<li>Store the packaged phagemid filtrate at 4 &#176;C. Note: Samples can be maintained for days to weeks without loss of activity.</li>
</ol>
3. Preparation of F+ Target Cells for Silencing
<ol>
	<li>Determine if the cells to be targeted for silencing express the F pilus14. If the F pilus is already present, proceed to step 4. 
	Note: Common laboratory strains of E. coli are annotated as F+ or F&#39; to indicate the presence of the F pilus in their genome or on a plasmid.</li>
	<li>Obtain an F+ strain of E. coli such as TOP10F&#39;. 
	Note: Ensure that the target strain carries a unique resistance marker in order to be separated from the F-plasmid donor after conjugation.</li>
	<li>To introduce F pilus by conjugation with a F+ strain, prepare 5 ml cultures of both the target strain and F-plasmid donor14. Grow the cells overnight at 37 &#176;C with shaking, in LB media with appropriate antibiotics.</li>
	<li>The following day, dilute both strains 1:100 in 5 ml of selective LB and continue culturing at 37 &#176;C with shaking.</li>
	<li>Determine the growth phase of the cells by measuring the optical density of the culture at 600 nm (OD600) using a benchtop spectrophotometer. Culture the cells for about 2 hr until an OD600 of 0.3 is attained, indicating log-phase growth15.</li>
	<li>Prepare 3 conjugation reactions in microcentrifuge tubes: 0.5 ml F-plasmid donor + 0.5 ml target strain, 0.5 ml F-plasmid donor + 0.5 ml LB media (negative control) and 0.5 ml target strain + 0.5 ml LB media (negative control). Allow the conjugation to proceed for 2 hr at 37 &#176;C with shaking.</li>
	<li>Plate 100 &#181;l of each conjugation reaction on selective LB agar with antibiotics specific to the F-plasmid (typically tetracycline) and target strain. Plate the negative control reactions to confirm that neither donor or recipient strain express both antibiotic resistances.</li>
</ol>
4.&#160;Infection with Packaged Phagemids for Silencing
<ol>
	<li>Inoculate a single colony of F+ target cells into a 5 ml culture of LB media with appropriate antibiotics. Incubate overnight at 37 &#176;C with shaking.</li>
	<li>The following day, dilute the F+ target cells 1:100 in 5 ml of selective LB media and continue to culture at 37 &#176;C with shaking.</li>
	<li>Determine the growth phase of the cells by measuring the optical density of the culture at 600 nm (OD600) using a benchtop spectrophotometer. Culture the cells for about 2 hr until an OD600 of 0.3 is attained, indicating log-phase growth15. Note: Expression of the F pilus and infection efficiency is highest in log phase.</li>
	<li>Add the M13-packaged phagemids (from step 2.6) to the target cells in a volume ratio of 1:100 to achieve nearly 99% infection of the target population. Allow the infection to proceed at 37 &#176;C with shaking for 30-60 min.</li>
	<li>Assay the sRNA-silencing phenotype according to the method of choice. 
	Note: For a fluorescent protein target, the silencing effect can be quantified directly by fluorometry10. Alternately, phenotypic assays may be used to observe the phenotypic consequences of gene knockdown8.</li>
	<li>Prepare a glycerol stock for the sRNA-expressing phagemid host following steps 1.2.14-1.2.16. Note: the phagemid will propagate in the host strain indefinitely and can be maintained with antibiotic similar to a conventional plasmid.</li>
</ol>

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The authors have nothing to disclose.

&#160;The present method achieved 80% reduction in mKate fluorescence levels compared to untargeted controls. This is in line with other RNA knockdown methods, where complete silencing is not observed and 50-90% efficiency is typical16,17. At the phenotypic level, CAT-targeted knockdowns were able to significantly attenuate chloramphenicol resistance, and eliminate it under some conditions.
The knockdown phenotype was detectable at the population level after only a few hours post-infection (Figure 3B). This illustrates an important feature of phage-based delivery: a high knockdown frequency can be obtained directly in batch culture without prior genetic modification. Unlike conventional genetic modifications using plasmid transformation or genomic integration, phage infection does not require that a population be re-grown from a single isolated colony. This allows the effects of phage infection to be explored in populations with complex spatial dynamics11, with pre-existing spatial structures like biofilms18, or in genetically mixed natural populations19.
A critical step in this method is the production of packaged phagemid at high titer. The metabolic burden associated with phage particle production may lead to high rates of mutation or plasmid loss in the phagemid production strain. It is recommended that the phagemid production strain be cultured directly from a single cotransformed colony and not refrigerated, frozen or sub-cultured prior to phage harvest. A low efficiency of co-transformation may also be observed when introducing the phagemid and helper plasmid to E. coli simultaneously. In this case, higher efficiencies may be obtained by transforming the helper plasmid first, then preparing competent cells bearing the helper plasmid for subsequent transformation with the phagemid.
Phagemid infection or sRNA expression also imposes a detectable metabolic burden on the target cells, and may result in some phenotypic perturbation. For example, a reduction in mKate2 fluorescence was observed even when cells were infected with a phagemid targeting CAT (Figure 3). Infection with M13 is not thought to trigger systemic stress responses in E. coli20, but may alter transcriptional patterns indirectly. Alternately, the GFP or ampicillin resistance markers included on the phagemid may compete for cellular resources, reducing mKate2 expression and growth9. Finally, the sRNA cassette itself may globally alter gene expression profiles by titrating the Hfq protein, or through off-target mRNA silencing. Off target effects are common in in vivo RNAi targeting21-23, but they have yet to be systematically investigated for this system.
One limitation of this method is that the infection efficiency is less than 100%, allowing some non-infected bacteria to persist in the population. The results of this work and earlier work10 suggest that noninfected cells represent 1-10% of the final population, and are responsible for most of the nonsilenced phenotypes observed. A variety of routes to M13-resistance are known, with the most common being mutational loss of pilus expression24. In light of these limitations, controls should be used to confirm high infection rates and knockdown efficiency.
Another potential limitation for some applications is the occasional transfer of contaminating helper phage. Although M13K07 contains a mutated packaging signal, it can be packaged into phage capsid at low frequency and transferred to infected populations, resulting in cells competent for phage production and the continued spread of phage beyond the initial infection event25. Modifications to the helper phage have proven effective at reducing nonspecific packaging, although sometimes at the cost of reduced phage production26.
Engineered bacteriophage have become an indispensable tool for E. coli synthetic biology, allowing quick delivery of new genes to a growing population. Recent work has produced intercellular communication circuits11 or expressed transcription factors to suppress antibiotic resistance pathways27. The protocol presented here adds to a growing collection of tools that allow control of bacterial physiology through programmed RNAs. CRISPR-Cas nucleases, when mutated to eliminate nuclease activity, have been shown to repress transcription at RNA-guided gene targets17,28. In contrast, sRNA silencing works at the translational level and does not require exogenous protein expression. Next-generation biotechnologies may unite transcriptional and translational control with phage-mediated delivery to program complex phenotypes in real time.

RNA-mediated gene knockdowns proceed in two stages. First, an RNA molecule is introduced to a cell line or organism of study. Second, endogenous RNA-binding proteins facilitate RNA-target recognition and produce the silencing effect. All RNA knockdown technologies benefit from the customizable nature of synthetic sRNAs, which can be easily produced to match a specific target of interest. However, the molecular details of RNA uptake and silencing vary widely across model system, constraining where and how RNA knockdowns can be applied.
In nematodes, double-stranded RNA (dsRNA) molecules can be introduced directly in the media or by feeding the worms with a population of dsRNA-expressing E. coli cells1,2. In Drosophila, RNAi can be achieved by microinjecting embryos with dsRNA3, or implemented in cell lines by simply adding dsRNA to the culture medium4. In mammalian cell lines, synthetic small interfering RNAs (siRNAs) may be delivered to living cells by electroporation1,2,5, packaged in liposomes3,6, or expressed from DNA plasmid vectors4,7. Once the RNA species reaches the cytosol, the RNAi pathway relies on the RISC complex to process dsRNA, facilitate antisense recognition of the target, and catalyze translational repression, mRNA degradation, or heterochromatin formation, depending on the host.
Because of these requirements, classical RNAi can be performed only in organisms that take up exogenous RNA efficiently and express RISC or a RISC-like activity. Notably, this excludes the model bacterium E. coli, which lacks the RNAi pathway. However, recent advances in synthetic biology provide the tools to solve both the delivery problem and the silencing problem.
In this protocol, sRNA constructs are expressed in E. coli from a DNA vector delivered to living cells using the M13 phagemid/helper system. A phagemid is any plasmid with a phage-derived f1 origin of replication. A helper plasmid, in this case M13KO, carries all the machinery required to produce viral particles, but is not itself competent for replication and packaging. When a phagemid and helper plasmid are co-transformed, the phagemid alone is replicated at the f1 origin, packaged and secreted. The vectorized phagemid is then competent to infect live E. coli via the F pilus.
In this system, the silencing effect is produced by custom sRNA cassettes combining a scaffold sequence with a target-binding sequence. The target-binding sequence is 24 base pairs antisense to an mRNA target, typically at the ribosome-binding site (RBS). The scaffold sequence, developed by Na and colleagues8, contains an Hfq-binding motif extracted from MicC, a small regulatory RNA endogenous to E. coli. The Hfq protein stimulates RNA-RNA binding and mRNA degradation, serving a role in this system similar to RISC in RNAi. Figure 1 depicts a complete scheme for phage-mediated sRNA knockdowns, including the sRNA cassette structure, phagemid vectorization, and silencing mechanism.
As a method to modulate gene expression in E. coli, sRNA silencing is simple, rapid and versatile. The targeted E. coli is not burdened beyond propagating the phagemid and expressing the sRNA. This may be relevant in the context of synthetic biology or basic research where the expression of larger heterologous constructs can strain cellular resources9. Phagemids with new targets can be produced with a single PCR and harvested one day after phagemid transformation. Finally, nearly any mRNA can be targeted. The sRNA regulation cassette (on a standard plasmid) has been shown to work on a variety of targets in metabolism with typical repression levels &#62;90%8.
This protocol reproduces and expands on earlier work using phagemid-born sRNA cassettes10. First, a packaged phagemid is introduced to a batch culture of E. coli cells and used to silence expression of the fluorescent protein mKate2. Subsequent fluorescence changes are monitored in real time. Second, knocking down the CAT gene is shown to reduce phenotypic chloramphenicol resistance on agar plates. In both cases, the phagemid itself carries a GFP marker, allowing infection rate to be measured independently of the knockdown efficiency.

<table><tbody><tr><td>Plasmid Miniprep Kit</td><td>Qiagen</td><td>27104</td><td></td></tr><tr><td>DpnI Enzyme</td><td>NEB</td><td>R0176S</td><td></td></tr><tr><td>Phusion High Fidelity Polymerase</td><td>NEB</td><td>M0530S</td><td></td></tr><tr><td>Taq 2x Master Mix</td><td>NEB</td><td>M0270L</td><td></td></tr><tr><td>M13KO7 Helper Phage</td><td>NEB</td><td>N0315S</td><td></td></tr><tr><td>DH5&amp;alpha; Competent Cells</td><td>Life Technologies</td><td>18265-017</td><td></td></tr><tr><td>TOP10F&#039; Cells</td><td>Life Technologies</td><td>C3030-03</td><td></td></tr><tr><td>LB Broth</td><td>Sigma</td><td>L3022-250G</td><td></td></tr><tr><td>Ampicillin</td><td>Sigma</td><td>A9393-5G</td><td></td></tr><tr><td>Kanamycin</td><td>Sigma</td><td>60615-5G</td><td></td></tr><tr><td>Chloramphenicol</td><td>Sigma</td><td>C0378-5G</td><td></td></tr><tr><td>Tetracycline</td><td>Sigma</td><td>87128-25G</td><td></td></tr></tbody></table>

phage-mediated delivery of targeted srna constructs to knock down gene expression in e. coli

RNA-mediated knockdowns are widely used to control gene expression. This versatile family of techniques makes use of short RNA (sRNA) that can be synthesized with any sequence and designed to complement any gene targeted for silencing. Because sRNA constructs can be introduced to many cell types directly or using a variety of vectors, gene expression can be repressed in living cells without laborious genetic modification. The most common RNA knockdown technology, RNA interference (RNAi), makes use of the endogenous RNA-induced silencing complex (RISC) to mediate sequence recognition and cleavage of the target mRNA. Applications of this technique are therefore limited to RISC-expressing organisms, primarily eukaryotes. Recently, a new generation of RNA biotechnologists have developed alternative mechanisms for controlling gene expression through RNA, and so made possible RNA-mediated gene knockdowns in bacteria. Here we describe a method for silencing gene expression in E. coli that functionally resembles RNAi. In this system a synthetic phagemid is designed to express sRNA, which may designed to target any sequence. The expression construct is delivered to a population of E. coli cells with non-lytic M13 phage, after which it is able to stably replicate as a plasmid. Antisense recognition and silencing of the target mRNA is mediated by the Hfq protein, endogenous to E. coli. This protocol includes methods for designing the antisense sRNA, constructing the phagemid vector, packaging the phagemid into M13 bacteriophage, preparing a live cell population for infection, and performing the infection itself. The fluorescent protein mKate2 and the antibiotic resistance gene chloramphenicol acetyltransferase (CAT) are targeted to generate representative data and to quantify knockdown effectiveness.

Silencing of mKate2 Fluorescence in Liquid Media
Figure 1 depicts the scheme for sRNA-mediated knockdowns described in this work, including the sRNA cassette design, phagemid vectorization, and silencing mechanism. Following protocol 1.2, the sRNA silencing cassette of plasmid pAB.001 was altered to target mKate. The sRNA cassette was synthetized and cloned into phagemid Litmus28i_J23115-B0032-GFP, a gift from Monica Ortiz and Drew Endy11. This phagemid carries markers for GFP expression and kanamycin resistance, allowing successful infections to be tracked. Packaged phagemid stocks were prepared following protocol 2.
A derivative of E. coli K12 MG1655 carrying a constitutively expressed, chromosomally integrated mKate2 marker was prepared for phage infection by conjugation with an F plasmid donor strain following protocol 3. The cells were grown to mid log phase and the phagemid introduced following protocol 4. After phagemid infection, 200 &#956;l cultures were transferred to a fluorescence plate reader and fluorescence was monitored continuously for 24 hr.
Figure 3 shows the effect of sRNA-mediated silencing on mKate2 expression. The strain infected with an anti-mKate2 phagemid showed no detectable mKate2 fluorescence over background. In contrast, this strain did express the GFP marker, indicating successful uptake of the phagemid. Uninfected control cells produced mKate2 fluorescence but not GFP. An additional control, in which the anti-mKate2 targeting domain was replaced with a sequence targeting CAT, had no effect on mKate2 fluorescence.
Silencing of Chloramphenicol Resistance on Agar Plates
Following protocol 1.1, an sRNA silencing cassette was produced to target CAT. A derivative of E. coli K12 MG1655 carrying a constitutively expressed, chromosomally integrated CAT marker was prepared for phage infection by conjugation with an F-plasmid donor strain following protocol 3. The cells were grown to mid log phase and the phagemid introduced following protocol 4. After 1 hr of incubation at 37 &#176;C, infected cells were serially diluted and plated at a range of chloramphenicol concentrations. Plates were incubated overnight, and the proportion of resistant cells at each chloramphenicol concentration was determined by counting colony forming units (CFUs) the next day.
Figure 4 shows the effect of sRNA-mediated silencing on the chloramphenicol resistance phenotype. Uninfected cells, or cells infected with phagemid targeting mKate2, were resistant to chloramphenicol at all concentrations tested. In contrast, cells infected with phagemid targeting CAT showed reduced survival at low chloramphenicol concentrations, and nearly 99% killing at higher concentrations. The addition of ampicillin, to select for only bacteria carrying the phagemid, reduced chloramphenicol survival to undetectable levels. This is consistent with earlier work showing that escape from phage infection is a common route to escape silencing10.
<img alt="Figure 1" src="/files/ftp_upload/53618/53618fig1.jpg" /> 
Figure 1: Gene Silencing in E. coli with sRNA Expression Cassettes Delivered by M13 Phage.&#160;The sRNA cassette is composed of 4 modules: the pR promoter (a constitutive promoter derived from bacteriophage lambda), a 24 bp targeting domain, an Hfq binding domain extracted from MicC and a transcriptional terminator8. The cassette is incorporated into a phagemid bearing the M13 f1 replication origin and packaged into the M13 capsid using the helper phage M13KO7. Packaged phagemids are able to infect E. coli expressing the F pilus, where sRNA expression begins. The sRNA then recruits the Hfq protein (depicted in red) and binds an antisense mRNA target near the ribosome binding site, resulting in translational repression and mRNA degradation. <a href="https://www.jove.com/files/ftp_upload/53618/53618fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53618/53618fig2.jpg" /> 
Figure 2: Primer Design and Site-Directed Mutagenesis of the sRNA Target Site. Two primers are designed with partial homology to an existing sRNA cassette. The forward primer contains 20 bp homologous to the pR promoter at the 5&#39; end, followed by 24 bp representing the new GUIDE sequence, then 18 bp homologous to the Hfq-binding domain at the 3&#39; end. The reverse primer is the exact reverse complement of the forward primer, with regions of homology to the existing sRNA cassette flanking the reverse complement of the new GUIDE sequence. Exact primer sequences are given in Table 3. Separate single-primer PCR reactions with the forward and the reverse primers produce linear, single-stranded DNA with the desired modified sequence. Annealing the forward and reverse reaction products, following clean-up as described in the protocol, results in double-stranded plasmid DNA bearing the desired modified sRNA cassette. <a href="https://www.jove.com/files/ftp_upload/53618/53618fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53618/53618fig3.jpg" /> 
Figure 3: Knockdown of a Chromosomally Integrated mKate2 Fluorescent Reporter. E. coli MG1655 K12 expressing mKate2 were either left untreated (red lines and bars), infected with an anti-mKate2 phagemid (green lines and bars), or infected with a control phagemid targeting CAT (blue lines and bars). (A) Untreated E. coli grew to higher saturation densities, indicating a metabolic cost to phage infection. Dotted lines indicate standard deviations of 3 replicates. (B) The mKate2 signal was reduced to near background levels in the anti-mKate2 treated strain, but not in control strains. (C) GFP fluorescence, also carried by the phagemid, was detectable only in phagemid-treated controls. (D, E) Final fluorescence readings after 24 hr of growth were normalized to OD. GFP signal, indicating phagemid infection, was absent in untreated controls, but also substantially reduced following anti-CAT phagemid treatment. This may indicate off-target effects of the phagemid. The mKate2 signal was reduced by anti-mKate2 phagemid treatment compared to untreated controls. The CAT-targeted control phagemid showed no effect on mKate2 fluorescence. Error bars represent standard deviation of 3 replicates. <a href="https://www.jove.com/files/ftp_upload/53618/53618fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53618/53618fig4.jpg" /> 
Figure 4: Knockdown of CAT Restores Chloramphenicol Sensitivity to a Genetically Resistant Population. E. coli MG1655 K12 expressing a chromosomally integrated CAT gene were left untreated (red bars), treated with a control phagemid targeting mKate2 (green bars), or treated with a phagemid expressing an anti-CAT sRNA (blue bars). After 1 hr of infection, viability on the indicated antibiotics was assessed by serial dilutions and plating. Strains treated with anti-CAT phagemid were significantly killed (&#62;90%) by chloramphenicol at higher concentrations, while control treatments were unaffected. Adding ampicillin to the culture plates positively selects for phagemid infection and eliminates uninfected cells. Under these conditions, no chloramphenicol resistant colonies were observed after anti-CAT treatment. This indicates that most survivors represent a failure of infection, rather than a failure of silencing. Error bars represent the standard deviation of 3 replicates. <a href="https://www.jove.com/files/ftp_upload/53618/53618fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>CAT TARGET sequence</td>
			<td>5&#39; - ATGGAGAAAAAAATCACTGGATAT - 3&#39;</td>
		</tr>
		<tr>
			<td>CAT GUIDE sequence</td>
			<td>5&#39; - ATATCCAGTGATTTTTTTCTCCAT - 3&#39;</td>
		</tr>
	</tbody>
</table>
Table 1: An example TARGET and GUIDE Sequence for the CAT Gene. Note the reverse complement relationship.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>pR promoter</td>
			<td colspan="5">TAACACCGTGCGTGTTGACTATTTTACCTCTGGCGGTGATAATGGTTGC</td>
		</tr>
		<tr>
			<td>GUIDE sequence</td>
			<td colspan="5">ATATCCAGTGATTTTTTTCTCCAT</td>
		</tr>
		<tr>
			<td>Hfq binding domain</td>
			<td colspan="5">TTTCTGTTGGGCCATTGCATTGCCACTGATTTTCCAACATATAAAAAGACAAGCCCGAACAGTCGTCCGGGCTTTTTT 
			TCTCGAG</td>
		</tr>
		<tr>
			<td>T1/TE terminator</td>
			<td colspan="5">CTCGAGCCAGGCATCAAATAAAACGAAAGGCTCAGTCGAAAGACTGGGCCTTTCGTTTTATCTGTTTTTGTCGGTGAA 
			CGCTCTCTACTAGAGTCACACTGGCTCACCTTCGGGTGGGCCTTTCTGCGTTTATA</td>
		</tr>
	</tbody>
</table>
Table 2: Sequence Components of the sRNA Cassette. Each sequence is written 5&#39;-3&#39;. The complete cassette is the concatenation of these 4 elements in order and comprises 292 bp.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Forward Primer</td>
			<td>5&#39; - CTGGCGGTGATAATGGTTGC[GUIDE]TTTCTGTTGGGCCATTGC - 3&#39;</td>
		</tr>
		<tr>
			<td>Reverse Primer</td>
			<td>5&#39; - GCAATGGCCCAACAGAAA[TARGET]GCAACCATTATCACCGCCAG - 3&#39;</td>
		</tr>
	</tbody>
</table>
Table 3: Primer Design to Alter Existing GUIDE Elements. The forward primer includes the last 20 bp of the pR promoter, the new GUIDE sequence, and the first 18 bp of the Hfq-binding domain. The TARGET sequence is the exact reverse complement of the GUIDE sequence. The reverse primer is the exact reverse complement of the forward primer.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Component</td>
			<td>Volume</td>
		</tr>
		<tr>
			<td>Template plasmid</td>
			<td>500 ng</td>
		</tr>
		<tr>
			<td>10 &#181;M Primer</td>
			<td>2.5 &#956;l</td>
		</tr>
		<tr>
			<td>10 mM dNTPs</td>
			<td>0.4 &#956;l</td>
		</tr>
		<tr>
			<td>High-fidelity Polymerase (2 U/&#956;l)</td>
			<td>0.5 &#956;l</td>
		</tr>
		<tr>
			<td>5x PCR Buffer</td>
			<td>10 &#956;l</td>
		</tr>
		<tr>
			<td>Total volume</td>
			<td>50 &#956;l</td>
		</tr>
	</tbody>
</table>
Table 4: Suggested Conditions for the Single-Primer Mutagenic PCR.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Step</td>
			<td>Temp</td>
			<td>Time</td>
		</tr>
		<tr>
			<td>Initial denaturation</td>
			<td>98 &#176;C</td>
			<td>30 sec</td>
		</tr>
		<tr>
			<td rowspan="3">30 cycles</td>
			<td>98 &#176;C</td>
			<td>10 sec</td>
		</tr>
		<tr>
			<td>55 &#176;C</td>
			<td>30 sec</td>
		</tr>
		<tr>
			<td>72 &#176;C</td>
			<td>120 sec</td>
		</tr>
		<tr>
			<td>Final Extension</td>
			<td>72 &#176;C</td>
			<td>300 sec</td>
		</tr>
		<tr>
			<td>Storage</td>
			<td>10 &#176;C</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 5: Suggested Thermocycler Protocol for the Single-Primer Mutagenic PCR.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Component</td>
			<td>Volume</td>
		</tr>
		<tr>
			<td>Template DNA</td>
			<td>1 &#956;l</td>
		</tr>
		<tr>
			<td>10 &#181;M Forward Primer</td>
			<td>0.5 &#956;l</td>
		</tr>
		<tr>
			<td>10 &#181;M Reverse Primer</td>
			<td>0.5 &#956;l</td>
		</tr>
		<tr>
			<td>Taq 2X Master Mix</td>
			<td>25 &#956;l</td>
		</tr>
		<tr>
			<td>Nuclease-free water</td>
			<td>23 &#956;l</td>
		</tr>
		<tr>
			<td>Total volume</td>
			<td>50 &#956;l</td>
		</tr>
	</tbody>
</table>
Table 6: Suggested Conditions for the Sequence Verification PCR.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Step</td>
			<td>Temp</td>
			<td>Time</td>
		</tr>
		<tr>
			<td>Initial denaturation</td>
			<td>95 &#176;C</td>
			<td>30 sec</td>
		</tr>
		<tr>
			<td rowspan="3">30 cycles</td>
			<td>95 &#176;C</td>
			<td>30 sec</td>
		</tr>
		<tr>
			<td>55 &#176;C</td>
			<td>30 sec</td>
		</tr>
		<tr>
			<td>68 &#176;C</td>
			<td>30 sec</td>
		</tr>
		<tr>
			<td>Final Extension</td>
			<td>68 &#176;C</td>
			<td>300 sec</td>
		</tr>
		<tr>
			<td>Storage</td>
			<td>10 &#176;C</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 7: Suggested Thermocycler Protocol for the Sequence Verification PCR.

Watch this Scientific Journal Video about Phage-mediated Delivery of Targeted sRNA Constructs to Knock Down Gene Expression in E. coli at JoVE.com

Phage-mediated Delivery of Targeted sRNA Constructs to Knock Down Gene Expression in E. coli

We describe a method to knock down gene expression in a growing population of E. coli cells using sequence-targeted sRNA expression cassettes delivered by an M13 phagemid vector.

Phage-mediated Delivery of Targeted sRNA Constructs to Knock Down Gene Expression in E. coli

RNA-mediated knockdowns are widely used to control gene expression. This versatile family of techniques makes use of short RNA (sRNA) that can be ...

phage-mediated-delivery-targeted-srna-constructs-to-knock-down-gene

Research

JoVE Journal

Biology

12.6K Views.  Institut National de la Santé et de la Recherche Médicale (INSERM), Université Paris Descartes. We describe a method to knock down gene expression in a growing population of E. coli cells using sequence-targeted sRNA expression cassettes delivered by an M13 phagemid vector.

Video: Phage-mediated Delivery of Targeted sRNA Constructs to Knock Down Gene Expression in E. coli

High efficiency, Site-specific Transfection of Adherent Cells with siRNA Using Microelectrode Arrays (MEA)

The discovery of RNAi pathway in eukaryotes and the subsequent development of RNAi agents, such as siRNA and shRNA, have achieved a potent method for silencing specific genes1-8 for functional genomics and therapeutics. A major challenge involved in RNAi based studies is the delivery of RNAi agents to targeted cells. Traditional non-viral delivery techniques, such as bulk electroporation and chemical transfection methods often lack the necessary spatial control over delivery and afford poor transfection efficiencies9-12. Recent advances in chemical transfection methods such as cationic lipids, cationic polymers and nanoparticles have resulted in highly enhanced transfection efficiencies13. However, these techniques still fail to offer precise spatial control over delivery that can immensely benefit miniaturized high-throughput technologies, single cell studies and investigation of cell-cell interactions. 
Recent technological advances in gene delivery have enabled high-throughput transfection of adherent cells14-23, a majority of which use microscale electroporation. Microscale electroporation offers precise spatio-temporal control over delivery (up to single cells) and has been shown to achieve high efficiencies19, 24-26. Additionally, electroporation based approaches do not require a prolonged period of incubation (typically 4 hours) with siRNA and DNA complexes as necessary in chemical based transfection methods and lead to direct entry of naked siRNA and DNA molecules into the cell cytoplasm. As a consequence gene expression can be achieved as early as six hours after transfection27. Our lab has previously demonstrated the use of microelectrode arrays (MEA) for site-specific transfection in adherent mammalian cell cultures17-19. In the MEA based approach, delivery of genetic payload is achieved via localized micro-scale electroporation of cells. An application of electric pulse to selected electrodes generates local electric field that leads to electroporation of cells present in the region of the stimulated electrodes. The independent control of the micro-electrodes provides spatial and temporal control over transfection and also enables multiple transfection based experiments to be performed on the same culture increasing the experimental throughput and reducing culture-to-culture variability. 
Here we describe the experimental setup and the protocol for targeted transfection of adherent HeLa cells with a fluorescently tagged scrambled sequence siRNA using electroporation. The same protocol can also be used for transfection of plasmid vectors. Additionally, the protocol described here can be easily extended to a variety of mammalian cell lines with minor modifications. Commercial availability of MEAs with both pre-defined and custom electrode patterns make this technique accessible to most research labs with basic cell culture equipment.

High efficiency, Site-specific Transfection of Adherent Cells with siRNA Using Microelectrode Arrays MEA

The article details the protocol for site-specific transfection of scrambled sequence of siRNA in an adherent mammalian cell culture using a microelectrode array (MEA).

The discovery of RNAi pathway in eukaryotes and the subsequent development of RNAi agents, such as siRNA and shRNA, have achieved a potent method for ...

Bioengineering

CRISPR Guide RNA Cloning for Mammalian Systems

The outlined protocol describes streamlined methods for the efficient and cost-effective generation of Cas9-associated guide RNAs. Two alternative strategies for guide RNA (gRNA) cloning are outlined based on the usage of the Type IIS restriction enzyme BsmBI in combination with a set of compatible vectors. Outside of the access to Sanger sequencing services to validate the generated vectors, no special equipment or reagents are required aside from those that are standard to modern molecular biology laboratories. The outlined method is primarily intended for cloning one single gRNA or one paired gRNA-expressing vector at a time. This procedure does not scale well for the generation of libraries containing thousands of gRNAs. For those purposes, alternative sources of oligonucleotide synthesis such as oligo-chip synthesis are recommended. Finally, while this protocol focuses on a set of mammalian vectors, the general strategy is plastic and is applicable to any organism if the appropriate gRNA vector is available.

Here, a simple, efficient, and cost-effective method of sgRNA cloning is outlined.

The outlined protocol describes streamlined methods for the efficient and cost-effective generation of Cas9-associated guide RNAs. Two alternative ...

Genetics

Producing Gene Deletions in Escherichia coli by P1 Transduction with Excisable Antibiotic Resistance Cassettes

A first approach to study the function of an unknown gene in bacteria is to create a knock-out of this gene. Here, we describe a robust and fast protocol for transferring gene deletion mutations from one Escherichia coli strain to another by using generalized transduction with the bacteriophage P1. This method requires that the mutation be selectable (e.g., based on gene disruptions using antibiotic cassette insertions). Such antibiotic cassettes can be mobilized from a donor strain and introduced into a recipient strain of interest to quickly and easily generate a gene deletion mutant. The antibiotic cassette can be designed to include flippase recognition sites that allow the excision of the cassette by a site-specific recombinase to produce a clean knock-out with only a ~100-base-pair-long scar sequence in the genome. We demonstrate the protocol by knocking out the tamA gene encoding an assembly factor involved in autotransporter biogenesis and test the effect of this knock-out on the biogenesis and function of two trimeric autotransporter adhesins. Though gene deletion by P1 transduction has its limitations, the ease and speed of its implementation make it an attractive alternative to other methods of gene deletion.

Producing Gene Deletions in Escherichia coli by P1 Transduction with Excisable Antibiotic Resistance Cassettes

Here we present a protocol for the use of pre-existing antibiotic resistance-cassette deletion constructs as a basis for making deletion mutants in other E. coli strains. Such deletion mutations can be mobilized and inserted into the corresponding locus of a recipient strain using P1 bacteriophage transduction.

A first approach to study the function of an unknown gene in bacteria is to create a knock-out of this gene. Here, we describe a robust and fast protocol ...

DNA Vector-based RNA Interference to Study Gene Function in Cancer

RNA interference (RNAi) inhibits gene expression by specifically degrading target mRNAs. Since the discovery of double-stranded small interference RNA (siRNA) in gene silencing1, RNAi has become a powerful research tool in gene function studies. Compared to genetic deletion, RNAi-mediated gene silencing possesses many advantages, such as the ease with which it is carried out and its suitability to most cell lines. Multiple studies have demonstrated the applications of RNAi technology in cancer research. In particular, the development of the DNA vector-based technology to produce small hairpin RNA (shRNA) driven by the U6 or H1 promoter has made long term and inducible gene silencing possible2,3. Its use in combination with genetically engineered viral vectors, such as lentivirus, facilitates high efficiencies of shRNA delivery and/or integration into genomic DNA for stable shRNA expression. 
We describe a detailed procedure using the DNA vector-based RNAi technology to determine gene function, including construction of lentiviral vectors expressing shRNA, lentivirus production and cell infection, and functional studies using a mouse xenograft model. 
Various strategies have been reported in generating shRNA constructs. The protocol described here employing PCR amplification and a 3-fragment ligation can be used to directly and efficiently generate shRNA-containing lentiviral constructs without leaving any extra nucleotide adjacent to a shRNA coding sequence. Since the shRNA-expression cassettes created by this strategy can be cut out by restriction enzymes, they can be easily moved to other vectors with different fluorescent or antibiotic markers. Most commercial transfection reagents can be used in lentivirus production. However, in this report, we provide an economic method using calcium phosphate precipitation that can achieve over 90% transfection efficiency in 293T cells. Compared to constitutive shRNA expression vectors, an inducible shRNA system is particularly suitable to knocking down a gene essential to cell proliferation. We demonstrate the gene silencing of Yin Yang 1 (YY1), a potential oncogene in breast cancer4,5, by a Tet-On inducible shRNA system and its effects on tumor formation. Research using lentivirus requires review and approval of a biosafety protocol by the Biosafety Committee of a researcher's institution. Research using animal models requires review and approval of an animal protocol by the Animal Care and Use Committee (ACUC) of a researcher's institution.

RNA interference (RNAi) possesses many advantages over gene knockout and has been broadly used as a tool in gene functional studies. The invention of DNA vector-based RNAi technology has made long term and inducible gene knockdown possible, and also increased the feasibility of gene silencing in vivo.

RNA interference (RNAi) inhibits gene expression by specifically degrading target mRNAs. Since the discovery of double-stranded small interference RNA ...

Medicine

Application of CRISPR Interference (CRISPRi) for Gene Silencing in Pathogenic Species of Leptospira

Leptospirosis is a global neglected zoonosis, responsible for at least 1 million cases per year and almost 60 thousand deaths. The disease is caused by pathogenic and virulent bacteria of the genus Leptospira, either by direct contact with the bacteria or indirectly by exposure to contaminated water or soil. Domestic and wild animals act as reservoir hosts of infection, shedding leptospires from colonized renal tubules of the kidney, via urine, into the environment. The generation of mutant strains of Leptospira is critical to evaluate and understand pathogenic mechanisms of infection. CRISPR interference (CRISPRi) has proven to be a straightforward, affordable, and specific tool for gene silencing in pathogenic Leptospira. Therefore, the methodological details of obtaining the plasmid constructs containing both dCas9 and guide RNA, delivery of plasmids to Leptospira by conjugation with the E. coli strain &#946;2163, and transconjugant recovery and evaluation, will be described. In addition, the recently described Hornsby-Alt-Nally (HAN) media allows for the relatively rapid isolation and selection of mutant colonies on agar plates.

Application of CRISPR Interference CRISPRi for Gene Silencing in Pathogenic Species of Leptospira

Here, the application of CRISPR interference (CRISPRi) for specific gene silencing in Leptospira species is described. Leptospira cells are transformed by conjugation with plasmids expressing dCas9 (catalytically "dead" Cas9) and a single-guide RNA (sgRNA), responsible for base pairing to the desired genomic target. Methods to validate gene silencing are presented.

Leptospirosis is a global neglected zoonosis, responsible for at least 1 million cases per year and almost 60 thousand deaths. The disease is caused by ...

Bacterial Delivery of RNAi Effectors: Transkingdom RNAi

RNA interference (RNAi) represents a high effective mechanism for specific inhibition of mRNA expression. Besides its potential as a powerful laboratory tool, the RNAi pathway appears to be promising for therapeutic utilization. For development of RNA interference (RNAi)-based therapies, delivery of RNAi-mediating agents to target cells is one of the major obstacles. A novel strategy to overcome this hurdle is transkingdom RNAi (tkRNAi). This technology uses non-pathogenic bacteria, e.g. Escherichia coli, to produce and deliver therapeutic short hairpin RNA (shRNA) into target cells to induce RNAi. A first-generation tkRNAi-mediating vector, TRIP, contains the bacteriophage T7 promoter for expression regulation of a therapeutic shRNA of interest. Furthermore, TRIP has the Inv locus from Yersinia pseudotuberculosis that encodes invasin, which permits natural noninvasive bacteria to enter &beta;1-integrin-positive mammalian cells and the HlyA gene from Listeria monocytogenes, which produces listeriolysin O. This enzyme allows the therapeutic shRNA to escape from entry vesicles within the cytoplasm of the target cell. TRIP constructs are introduced into a competent non-pathogenic Escherichia coli strain, which encodes T7 RNA polymerase necessary for the T7 promoter-driven synthesis of shRNAs. A well-characterized cancer-associated target molecule for different RNAi strategies is ABCB1 (MDR1/P-glycoprotein, MDR1/P-gp). This ABC-transporter acts as a drug extrusion pump and mediates the "classical" ABCB1-mediated multidrug resistance (MDR) phenotype of human cancer cells which is characterized by a specific cross resistance pattern. Different ABCB1-expressing MDR cancer cells were treated with anti-ABCB1 shRNA expression vector bearing E. coli. This procedure resulted in activation of the RNAi pathways within the cancer cells and a considerable down regulation of the ABCB1 encoding mRNA as well as the corresponding drug extrusion pump. Accordingly, drug accumulation was enhanced in the pristine drug-resistant cancer cells and the MDR phenotype was reversed. By means of this model the data provide the proof-of-concept that tkRNAi is suitable for modulation of cancer-associated factors, e.g. ABCB1, in human cancer cells.

For development of RNA interference (RNAi)-based therapies, a novel strategy was developed, transkingdom RNAi (tkRNAi). This technology uses non-pathogenic bacteria to produce and deliver therapeutic short hairpin RNA (shRNA) into target cells. Here, tkRNAi was successfully applied for reversal of classical ABCB1-mediated multidrug resistance (MDR) of cancer cells.

RNA interference (RNAi) represents a high effective mechanism for specific inhibition of mRNA expression. Besides its potential as a powerful laboratory ...

In Vitro Selection of Engineered Transcriptional Repressors for Targeted Epigenetic Silencing

Gene inactivation is instrumental to study gene function and represents a promising strategy for the treatment of a broad range of diseases. Among traditional technologies, RNA interference suffers from partial target abrogation and the requirement for life-long treatments. In contrast, artificial nucleases can impose stable gene inactivation through induction of a DNA double strand break (DSB), but recent studies are questioning the safety of this approach. Targeted epigenetic editing via engineered transcriptional repressors (ETRs) may represent a solution, as a single administration of specific ETR combinations can lead to durable silencing without inducing DNA breaks.
ETRs are proteins containing a programmable DNA-binding domain (DBD) and effectors from naturally occurring transcriptional repressors. Specifically, a combination of three ETRs equipped with the KRAB domain of human ZNF10, the catalytic domain of human DNMT3A and human DNMT3L, was shown to induce heritable repressive epigenetic states on the ETR-target gene. The hit-and-run nature of this platform, the lack of impact on the DNA sequence of the target, and the possibility to revert to the repressive state by DNA demethylation on demand, make epigenetic silencing a game-changing tool. A critical step is the identification of the proper ETRs&#39; position on the target gene to maximize on-target and minimize off-target silencing. Performing this step in the final ex vivo or in vivo preclinical setting can be cumbersome.
Taking the CRISPR/catalytically dead Cas9 system as a paradigmatic DBD for ETRs, this paper describes a protocol consisting of the in vitro screen of guide RNAs (gRNAs) coupled to the triple-ETR combination for efficient on-target silencing, followed by evaluation of the genome-wide specificity profile of top hits. This allows for reduction of the initial repertoire of candidate gRNAs to a short list of promising ones, whose complexity is suitable for their final&#160;evaluation in the therapeutically relevant setting of interest.

In Vitro Selection of Engineered Transcriptional Repressors for Targeted Epigenetic Silencing

Here, we present a protocol for the in vitro selection of engineered transcriptional repressors (ETRs) with high, long-term, stable, on-target silencing efficiency and low genome-wide, off-target activity. This workflow allows for reducing an initial, complex repertoire of candidate ETRs to a short list, suitable for further evaluation in therapeutically relevant settings.

Gene inactivation is instrumental to study gene function and represents a promising strategy for the treatment of a broad range of diseases. Among ...

Effective Oral RNA Interference (RNAi) Administration to Adult Anopheles gambiae Mosquitoes

RNA interference has been a heavily utilized tool for reverse genetic analysis for two decades. In adult mosquitoes, double-stranded RNA (dsRNA) administration has been accomplished primarily via injection, which requires significant time and is not suitable for field applications. To overcome these limitations, here we present a more efficient method for robust activation of RNAi by oral delivery of dsRNA to adult Anopheles gambiae. Long dsRNAs were produced in Escherichia coli strain HT115 (DE3), and a concentrated suspension of heat-killed dsRNA-containing bacteria in 10% sucrose was offered on cotton balls ad-libitum to adult mosquitoes. Cotton balls were replaced every 2 days for the duration of the treatment. Use of this method to target doublesex (a gene involved in sex differentiation) or fork head (which encodes a salivary gland transcription factor) resulted in reduced target gene expression and/or protein immunofluorescence signal, as measured by quantitative Real-Time PCR (qRT-PCR) or fluorescence confocal microscopy, respectively. Defects in salivary gland morphology were also observed. This highly flexible, user-friendly, low-cost, time-efficient method of dsRNA delivery could be broadly applicable to target genes important for insect vector physiology and beyond.

Effective Oral RNA Interference RNAi Administration to Adult Anopheles gambiae Mosquitoes

The oral administration of dsRNA produced by bacteria, a delivery method for RNA interference (RNAi) that is routinely used in Caenorhabditis elegans, was successfully applied here to adult mosquitoes. Our method allows for robust reverse genetics studies and transmission-blocking vector studies without the use of injection.

RNA interference has been a heavily utilized tool for reverse genetic analysis for two decades. In adult mosquitoes, double-stranded RNA (dsRNA) ...

Phage Transduction: A Method to Transfer Ampicillin Resistance from Donor to Recipient E. coli

Source: Alexander S. Gold1, Tonya M. Colpitts1 
1 Department of Microbiology, Boston University School of Medicine, National Emerging Infections Diseases Laboratories, Boston, MA
Transduction is a form of genetic exchange between bacteria that utilizes bacteriophages, or phages, a class of virus that infects exclusively prokaryotic organisms. This form of DNA transfer, from one bacterium to another by way of a phage, was discovered in 1951 by Norton Zinder and Joshua Ledererg, who termed the process &#34;transduction&#34; (1). Bacteriophages were first discovered in 1915 by British bacteriologist Frederick Twort, then independently discovered again in 1917 by French-Canadian microbiologist Felix d&#39;Herelle (2). Since then, the structure and function of these phages have been widely characterized (3), dividing these phages into two classes. The first of these classes are the lytic phages which upon infection multiply within the host bacterium, disrupting the bacterial metabolism, lysing the cell, and releasing progeny phage (4). As a result of this anti-bacterial activity and the increasing prevalence of antibiotic-resistant bacteria, these lytic phages have recently proved useful as a substitute treatment for antibiotics. The second of these classes are the lysogenic phages which can either multiply within the host via the lytic cycle or enter a quiescent state in which their genome is integrated into that of the host (Figure 1), a process known as lysogeny, with the ability for phage production to be induced in multiple later generations (4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10517/10517fig1.jpg" /> 
Figure 1: Infection of host cell by bacteriophage. Adsorption by the phage to the bacterial cell wall via interactions between the tail fibers and receptor (purple). Once on the cell surface, the phage is irreversibly attached to the bacterial cell using the base plate (black) which is moved to the cell wall by the contractile sheath (yellow). Phage genome (red) then enters the cell and integrates into the host cell genome.
While bacterial transduction is a naturally occurring process, using modern technology it has been manipulated for the transfer of genes into bacteria in the laboratory setting. By inserting genes of interest into the genome of a lysogenic phage, such as phage, one is able to transfer these genes into the genomes of bacteria and consequently express them within these cells. While other methods of gene transfer, such as transformation, use a plasmid for gene transfer and expression, the insertion of the phage genome into that of the recipient bacterium not only has the potential to confer new traits to this bacterium, but also allows for naturally occurring mutations and other factors of the cellular environment to alter the function of the transferred gene.
Compared to other methods of horizontal gene transfer, such as conjugation, transduction is fairly flexible in the criteria required for donor and recipient cells. Any genetic element that can fit inside the genome of the phage being used can be transferred from any strain of donor bacteria to any strain of recipient bacteria as long as both are permissive to the phage, requiring the expression of necessary phage receptors on the cell surfaces. Once this gene is moved out of the donor genome and packaged into the phage, it can be transferred to the recipient. Following transduction, it is necessary to select for recipient bacteria that contain the gene of interest have to be selected for. This could be done by use of a genetic marker, such as a FLAG-tag or polyhistidine-tag, to mark the gene of interest, or the intrinsic function of the gene, in the case of genes that encode for antibiotic resistance. Additionally, PCR could be used to further confirm successful transduction. By using primers for a region within the gene of interest and comparing signal to a positive control, bacteria that has the gene of interest, and a negative control, bacteria that underwent the same steps as the transduction reaction with no phage. While bacterial transduction is a useful tool in molecular biology, it has and continues to play an important role in the evolution of bacteria, particularly with regard to the recent rise of antibiotic resistance.
In this experiment, bacterial transduction was used to transfer the gene encoding for resistance to the antibiotic ampicillin from the W3110 strain of E.coli to the J53 strain via the P1 bacteriophage (5). This experiment consisted of two main steps. First, the preparation of P1 phage containing the ampicillin resistance gene from the donor strain. Second, the transfer of this gene to the recipient strain by transduction with the P1 phage (Figure 1). Once carried out, the successful transfer of the ampicillin resistance gene could be determined by qPCR (Figure 2). If transduction was successful, the J53 strain of E. coli would be resistant to ampicillin, and the gene conferring this resistance detectable by qPCR. If unsuccessful, there would be no detection of the ampicillin resistance gene and ampicillin would still function as an effective antibiotic against the J53 strain.

<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/10517/10517fig2.jpg" /> 
Figure 2: The confirmation of successful transduction by qPCR. By comparing the Cq values detected for the gene of interest from the transduction reaction and the negative control reaction, and normalizing these values against a housekeeping gene, one was able to confirm that bacterial transduction was successful.

Phage Transduction: Transferring Ampicillin Resistance in E. coli

Source: Alexander S. Gold1, Tonya M. Colpitts1
1 Department of Microbiology, Boston University School of Medicine, National Emerging Infections Diseases ...

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RNAi Plating for C. elegans Feeding: A Technique to Induce Target dsRNA Expression in E. coli

Source: Kolundzic, E., et al. <a href="https://www.jove.com/video/56889/application-rnai-heat-shock-induced-transcription-factor-expression">Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in&#160;C. elegans</a>.&#160;J. Vis. Exp.&#160;(2018).
This video describes the principles behind RNAi treatment in C. elegans and demonstrates a protocol to knockdown lin-35 in a transgenic worm strain.

Source: Kolundzic, E., et al. Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in&#160;C. ...

Phage-mediated Delivery of Targeted sRNA Constructs to Knock Down Gene Expression in E. coli

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