This work was supported by Consejo Nacional de Ciencia y Tecnolog&#237;a, CONACYT, M&#233;xico (grant number 154833) and Universidad Nacional Aut&#243;noma de M&#233;xico. The authors wish to thank V&#237;ctor Gonz&#225;lez, Rosa I. Santamar&#237;a, Patricia Bustos, and Soledad Ju&#225;rez for their administrative and technical advice.

1. CAPRRESI Protocol
NOTE: Figure 1 represents the overall CAPRRESI protocol. This technique is based on an in silico design and the subsequent construction of the desired chimeras.
<ol>
	<li>Selection of the cloning plasmid, pUC18 or pUC19.
	<ol>
		<li>Select the pUC vector that best fits technical demands. 
		NOTE: Both vectors have the same sequence, except for the orientation of the multiple cloning site. This is important for the design of oligonucleotides and the PCR plasmid disruption.</li>
	</ol>
	</li>
	<li>In silico analysis of the wildtype genes and pUC18/19 sequences.
	<ol>
		<li>Obtain the DNA sequences of the wildtype genes and save them in a FASTA format file (e.g., genes.fas). 
		NOTE: The sequences of pUC18/19 vectors are contained in the pUC.fas file (Figure 1, step 1).</li>
		<li>Determine which restriction enzymes do not cut the wildtype genes and pUC18/19 sequences by running the REsearch.pl script. Alternatively, perform a restriction enzyme site search with an online tool (Figure 1, step 2).
		<ol>
			<li>Type the following into a terminal: 
			perl REsearch.pl genes.fas 
			perl REsearch.pl pUC.fas 
			NOTE: These commands will create the following output files: genes_lin.fas, genes_re.fas, pUC_lin.fas, and pUC_re.fas.</li>
			<li>Select the appropriate restriction enzymes for cloning the wildtype genes into the multiple cloning site of the chosen pUC18/19 vector by comparing the genes_re.fas and pUC.fas files. Choose the orientation of the insert relative to the ampicillin resistance gene (ampR) of the pUC18/19 vector.</li>
		</ol>
		</li>
		<li>Design forward and reverse oligonucleotides to amplify the coding region of the wildtype genes (external oligonucleotides). Include the proper restriction enzyme site at each 5&#39; end of the oligonucleotides; this will define the insert orientation.
		<ol>
			<li>Include a functional ribosome binding site in the forward oligonucleotides.</li>
			<li>Insert both wildtype genes in the same orientation inside the vector.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Selection of the breaking regions for chimera assembly and plasmid disruption.
	<ol>
		<li>Obtain the amino acid sequence of the wildtype and the ampR genes by running the translate.pl script (Figure 1, step 3).
		<ol>
			<li>Type the following into a terminal: 
			perl translate.pl genes.fas 
			perl translate.pl ampR.fas 
			NOTE: This script will create the following output files: genes_aa.fas and ampR_aa.fas.</li>
		</ol>
		</li>
		<li>Globally align the two wildtype amino acid sequences with an appropriate software (e.g., MUSCLE)9.</li>
		<li>Select the desired break regions (3-6 amino acids) for chimera assembly based on the sequence alignment. Save them into a file in FASTA format (e.g., regions.fas).</li>
		<li>Locate the DNA sequences that code for the selected amino acid stretches on both wildtype genes.</li>
	</ol>
	</li>
	<li>In silico generation of synonym DNA sequences containing restriction enzyme sites.
	<ol>
		<li>Obtain all the synonym DNA sequences that code for each amino acid sequence stretch selected as breaking regions (Figure 1, step 4); these sequences were stored in the regions.fas file.
		<ol>
			<li>Type the following into a terminal: 
			perl synonym.pl regions.fas 
			NOTE: This script will create the regions_syn.fas output file.</li>
		</ol>
		</li>
		<li>Search for restriction enzyme sites found on synonym DNA sequences.
		<ol>
			<li>Type the following into a terminal: perl REsynonym.pl regions_syn.fas 
			NOTE: This script will create the regions_syn-re.fas output file.</li>
		</ol>
		</li>
		<li>Choose one restriction enzyme that is shared between synonym sequences of both wildtype genes; this site will be used to assemble chimeras via restriction enzyme digestion. Verify that the chosen restriction enzyme cut neither the wildtype genes nor the pUC18/19 vector sequences.
		<ol>
			<li>Compare the restriction enzymes found on the genes_re.fas, pUC_re.fas, and regions_syn-re.fas files.</li>
		</ol>
		</li>
		<li>In silico substitute the originals for their synonym DNA sequences at the corresponding loci on both wildtype genes. Append synonym DNA sequences into the wildtype sequence file (genes.fas).</li>
		<li>Get the amino acid sequence of genes contained on the genes.fas file (perl translate.pl genes.fas).</li>
		<li>Align the amino acid sequences translated from wildtype and synonym DNA sequences. Verify that both sequences are the same.</li>
		<li>Repeat all the steps of this section with the ampR gene present on the pUC18/19 vector. 
		NOTE: The goal is to disrupt the ampR gene (file ampR.fas) into two complementary parts. The restriction enzyme selected to disrupt the ampR gene should be different from the one used for chimera assembly.</li>
	</ol>
	</li>
	<li>Independent cloning of the two wildtype genes into the selected pUC18/19 vector (Figure 1, step 3).
	<ol>
		<li>Purify the chosen pUC18/19 plasmid DNA using a plasmid DNA purification kit.
		<ol>
			<li>For example, transform E. coli DH5&#945; with pUC18 plasmid and grow the transformants overnight at 37 &#176;C on solid LB-0.3 mg/mL ampicillin (Amp). Pick a transformant colony, inoculate a new LB-0.3 mg/mL Amp plate, and grow it overnight at 37 &#176;C.</li>
			<li>Take part of the previous culture and grow it in liquid LB-0.3 mg/mL Amp for 6-8 h at 37 &#176;C. Extract the plasmid DNA using a kit.</li>
		</ol>
		</li>
		<li>PCR amplify wildtype genes from the total genomic DNA using the appropriate external oligonucleotides. Use a high-fidelity DNA polymerase if possible. Select the DNA polymerase that maximizes the yield. Perform PCR according to the manufacturer&#39;s guidelines and the melting temperature of the oligonucleotides.
		<ol>
			<li>Run PCR cycles, depending upon the DNA polymerase, amplicon size, template, and oligonucleotides used. For example, to amplify rpoD DNA, prepare a 50 &#181;L reaction: 5 &#181;L of 10x buffer, 2 &#181;L of 50 mM MgSO4, 1 &#181;L of 10 mM dNTP mix, 2 &#181;L of each 10-&#181;M oligonucleotide (Table 2), 2 &#181;L of template DNA (E. coli DH5&#945; total DNA), 35.8 &#181;L of ultra-pure water, and 0.2 &#181;L of high-fidelity DNA polymerase. Run the following PCR cycles: 1 min at 94 &#176;C for the initial denaturation followed by 30 cycles of amplification (30 s at 94 &#176;C to denature, 30 s at 60 &#176;C to anneal the oligonucleotides, and 2 min at 68 &#176;C to extend).</li>
			<li>Dissolve 1.2 g of agarose in 100 mL of double-distilled water by heating them in the microwave. Assemble a gel tray and comb and put them into the horizontal gel caster. Fill the gel tray with the melted agarose solution and let it solidify. Remove the comb.</li>
			<li>Place the gel into the electrophoresis chamber. Fill chamber with 1x Tris-acetate-EDTA buffer. Load 50 &#181;L of PCR products into the gel wells. Electrophorese the samples (e.g., at 110 V for 1 h).</li>
			<li>After electrophoresis, stain the gel in 100 mL of double-distilled water containing 100 &#181;L of 1 mg/mL ethidium bromide solution for 10 min with slow shaking. Wash the gel in double-distilled water for another 10 min in slow shaking; use a horizontal shaker. 
			Caution: Ethidium bromide is a toxic compound; wear protective gloves and a cotton laboratory coat.</li>
			<li>Purify wildtype gene DNA from the corresponding bands of the gel using a kit. Visualize the bands by placing the stained gel in a UV chamber (recommended wavelength: 300 nm). Keep the exposure of the gel to a minimum. Use a DNA purification kit and follow the manufacturer&#39;s instructions. 
			Caution: UV radiation is dangerous; wear a protective shield, glasses, and a cotton laboratory coat.</li>
		</ol>
		</li>
		<li>Digest purified wildtype genes and pUC18/19 plasmid DNA with the restriction enzymes according to the manufacturer&#39;s instructions. Use fast digestion enzymes when possible. For example, digest 6 &#181;L of pUC18 DNA (300 ng/&#181;L) in 10 &#181;L of ultrapure water, 2 &#181;L of 10X buffer, 1 &#181;L of KpnI restriction enzyme, and 1 &#181;L of XbaI restriction enzyme. Leave the reaction at 37 &#176;C for 30 min.
		<ol>
			<li>Inactivate the restriction enzymes according to the manufacturer&#39;s guidelines. For example, inactivate the double-digestion reaction KpnI-XbaI at 80 &#176;C for 5 min.</li>
		</ol>
		</li>
		<li>Mix insert:vector DNA at a volumetric ratio of 3:1. Follow the manufacturer&#39;s instructions for a ligation reaction. Use fast ligation enzymes when possible (leave the reaction at 25 &#176;C for 10 min). 
		NOTE: Typically, the DNA concentration after purification using the kits is good enough for digestion and ligation reactions. For example, add 6 &#181;L of rpoD DNA (KpnI-XbaI, 150 ng/&#181;L), 3 &#181;L of pUC18 DNA (KpnI-XbaI, 150 ng/&#181;L), 10 &#181;L of 2x buffer, 1 &#181;L of ultrapure water, and 1 &#181;L of T4 DNA ligase. Leave ligation reaction at 25 &#176;C for 10 min.</li>
		<li>Transform the E. coli DH5&#945; with 2-4 &#181;L of the ligation reaction, as described in the Supplementary Materials. Plate the transformed cells into solid LB/Amp/X-gal/IPTG plates. Leave the plates overnight at 37 &#176;C.</li>
		<li>Select white-colored transformant E. coli colonies and streak them on a new LB/Amp plate. Leave the plates overnight at 37 &#176;C.</li>
		<li>Perform a colony PCR to choose candidates for sequencing the reaction, as described in the Supplementary Materials. Extract plasmid DNA from candidates. Alternatively, digest candidate plasmid DNA with the same restriction enzymes used for cloning the inserts.</li>
		<li>Sequence candidate constructions with a sequencing service provider10. Assemble the sequence in silico using a DNA assembler11(Figure 1, step 5).</li>
	</ol>
	</li>
	<li>Plasmid disruption by PCR followed by restriction enzyme digestion.
	<ol>
		<li>Design in silico forward and reverse oligonucleotides at break regions for wildtype and ampR genes, respectively. Substitute in silico the wildtype fragment for its corresponding synonym DNA sequence (obtained in step 1.4). 
		NOTE: This synonym DNA sequence contains a restriction enzyme site. Forward and reverse oligonucleotides overlap at the restriction enzyme site. Oligonucleotides should range from 21-27 nucleotides, end with a cytosine or guanine, and contain a restriction enzyme site. A forward oligonucleotide has the same sequence as its target region on the template DNA. A reverse oligonucleotide is the reverse complementary sequence of the target region on the template DNA.</li>
		<li>Use the two wildtype gene constructions as the DNA template for PCR reactions (e.g., pUC18rpoD and pUC18sigA). Obtain two complementary parts of each construction (pUC18rpoD&#963;1-&#963;2, pUC18rpoD&#963;3-&#963;4, pUC18sigA&#963;1-&#963;2, and pUC18sigA&#963;3-&#963;4) (Figure 1, step 6).</li>
		<li>Load DNA samples into an agarose gel 1-1.2% [w/v]. Separate the PCR products from the DNA template by electrophoresis (e.g., 110 V for 1 h). Alternatively, digest the PCR reactions with DpnI to break the DNA template. 
		NOTE: DpnI enzyme recognizes only methylated DNA sequences (5&#39;-GATC-3&#39;).
		<ol>
			<li>Purify the samples using a DNA purification kit. Follow the manufacturer&#39;s guidelines.</li>
		</ol>
		</li>
		<li>Double-digest all the complementary DNA fragments with the appropriate restriction enzymes according to the manufacturer&#39;s directions. Use fast-digest restriction enzymes when possible. For example, double-digest the pUC18rpoD&#963;1-&#963;2 DNA fragment with AflII and SpeI using the manufacturer&#39;s protocol. Leave the digestion reaction at 37 &#176;C for 15 min.</li>
		<li>Inactivate the restriction enzymes according to the manufacturer&#39;s guidelines. For example, inactivate AflII-SpeI at 80 &#176;C for 20 min.</li>
	</ol>
	</li>
	<li>Plasmid recovery using DNA ligation and bacterial transformation.
	<ol>
		<li>Mix the proper DNA fragments in a volumetric 1:1 ratio to assemble the desired chimeric gene (Figure 1, step 7). 
		NOTE: In this way, the integrity of the ampR gene is restored, producing a functional protein.
		<ol>
			<li>For example, mix 4 &#181;L of pUC18rpoD&#963;1-&#963;2 DNA (digested with AflII-SpeI, 150 ng/&#181;L), 4 &#181;L of pUC18sigA&#963;3-&#963;4 DNA (AflII-SpeI, 150 ng/&#181;L), 10 &#181;L of 2x buffer, 2 &#181;L of ultrapure water, and 1 &#181;L of T4 DNA ligase; the DNA concentration after kit purification is good enough for digestion and subsequent ligation. Leave the ligation reaction at 25 &#176;C for 10 min.</li>
		</ol>
		</li>
		<li>Transform E. coli DH5&#945; with 3-5 &#181;L of the chimera assembly ligation reaction (Supplementary Materials). Grow the transformant cells in LB/Amp/X-gal/IPTG plates overnight at 37 &#176;C overnight.</li>
		<li>Select white-colored transformant E. coli colonies and streak them on a new LB/Amp plate. Leave the plates overnight at 37 &#176;C.</li>
		<li>Perform a colony PCR to choose candidates for a sequencing reaction, as described in the Supplementary Materials. Visualize bands in a 1-1.2% (w/v) agarose gel by performing electrophoresis (described in step 2.3.2.1).</li>
		<li>Grow cultures from positive candidates. Extract plasmid DNA from them using a plasmid DNA purification kit.</li>
	</ol>
	</li>
</ol>
2. Sequence Candidate Chimeric Constructions
<ol>
	<li>Make sure the quality of the plasmid DNA is good enough for the sequencing reaction by following guidelines from the sequence service provider. Extract DNA using purification kits. For example, on the internet page of the sequence service provider, pay special attention to the recommended DNA concentration for the samples. Obtain the concentration of samples using a DNA quantification instrument.</li>
	<li>Sequence candidate chimeric constructions with a sequencing service provider10.</li>
	<li>Assemble sequencing reads with an in silico DNA assembler11.</li>
	<li>Align assembled versus in silico-designed chimeric sequences using sequence alignment tools9,12. Verify that the chimeric gene was fused successfully.</li>
</ol>
3. Making Preparations
NOTE: All steps involving living cells should be performed in a clean laminar flow hood with the Bunsen burner on.
<ol>
	<li>Producing E. coli DH5&#945;-competent cells.
	<ol>
		<li>To produce E. coli-competent cells, follow published protocols13 or see the Supplementary Materials. Alternatively, use commercially available competent cells.</li>
	</ol>
	</li>
	<li>Transforming E. coli DH5&#945;-competent cells.
	<ol>
		<li>For the chemical transformation of E. coli cultures, follow published protocols13 or see the Supplementary Materials. Alternatively, use electroporation for bacterial transformation. If commercially available competent cells are used, follow the manufacturer&#39;s guidelines.</li>
	</ol>
	</li>
	<li>Purifying genomic and plasmid DNA from E. coli DH5&#945;.
	<ol>
		<li>Perform nucleic acid extraction, as stated in the Supplementary Materials, using commercially available kits or following published protocols13.</li>
	</ol>
	</li>
	<li>Perform colony PCR.
	<ol>
		<li>Use this technique to identify candidate transformant colonies for subsequent sequencing reaction. 
		NOTE: This method does not represent a final verification of the integrity of the sequences. A detailed description of this technique is available in the Supplementary Materials.</li>
	</ol>
	</li>
	<li>Preparing the solutions.
	<ol>
		<li>See Table 1 for more information about solution preparation.</li>
	</ol>
	</li>
	<li>Download the scripts and sequence files required for the CAPRRESI protocol.
	<ol>
		<li>Download gene bank files containing the complete genome sequence of the desired species, extract the DNA sequence of the chosen gene using a genome browser, and save it in fasta file format. Download the Perl scripts required for the CAPRRESI protocol. Store the scripts and the sequence files in the same directory.</li>
	</ol>
	</li>
</ol>

<ol>
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The authors declare that they have no competing financial interests.

CAPRRESI was designed as an alternative to the PRM2. The original PRM is a powerful technique; it allows for the fusion of DNA sequences along any part of the selected genes. For PRM, wildtype genes should be cloned into the same plasmid. After that, oligonucleotides are designed inside wildtype and antibiotics resistance genes found on the plasmid. Plasmid disruption is achieved by PCR using blunt-ended, high-fidelity DNA polymerases and previously designed oligonucleotides. The ligation of compl...

Chimeric gene assembly has been widely used in molecular biology to elucidate protein function and/or for biotechnological purposes. Different methods exist for fusing genes, such as overlapping PCR product amplification1, plasmid recovery2, homologous recombination3, CRISPR-Cas9 systems4, site-directed recombination5, and Gibson assembly6. Each of these offers different technical advantages; for example, the flexibility of overlapping PCR design, the in vivo selection of constructions during plasmid recovery, or the high efficiency of CRISPR-Cas9 and Gibson systems. On the other hand, some difficulties can arise while performing some of these methods; for example, the first two approaches rely on blunt-ended DNA fragments, and ligation of these types of products could be technically challenging compared to sticky-ended ligation. Site-directed recombination can leave traces of extra DNA sequences (scars) on the original, like in the Cre-loxP system5. CRISPR-Cas9 can sometimes modify other genome regions in addition to the target site4.
Here, we introduce chimera assembly by plasmid recovery and restriction enzyme site insertion (CAPRRESI), a protocol for fusing protein-coding genes that combines the plasmid recovery method (PRM) with the insertion of restriction enzyme sites on synonym DNA sequences, enhancing ligation efficiency. To ensure amino acid sequence integrity, restriction enzyme sites are inserted on synonym DNA sequence stretches. Among the benefits of CAPPRESI are that it can be performed using ordinary laboratory reagents/tools (e.g., enzymes, competent cells, solutions, and thermocycler) and that it can give quick results (when the appropriate enzymes are used). Relying on restriction enzyme sites that emerge from synonym DNA sequences can limit the selection of the exact fusion points inside the proteins of interest. In such cases, target genes should be fused using overlapping oligonucleotides, and restriction enzyme sites should be inserted onto the resistance gene of the vector.
CAPRRESI consists of seven simple steps (Figure 1): 1) selection of the cloning vector, pUC18 or pUC196; 2) in silico analysis of the wildtype sequences to be fused; 3) selection of breaking regions for chimera assembly and plasmid disruption; 4) in silico generation of synonym DNA sequences containing restriction enzyme sites; 5) independent cloning of the wildtype genes into the selected plasmid; 6) plasmid disruption by PCR, followed by restriction enzyme digestion; and 7) plasmid recovery using DNA ligation and bacterial transformation. Chimeric genes produced by this technique should be verified with sequencing.
The pUC18/19 vectors offer technical advantages for cloning and chimera assembly, such as small size (i.e., 2,686 base pairs), high copy number, advantageous sequencing reaction features, and commercial availability7. Here, an Escherichia coli host was used to assemble and handle the chimeras because bacterial cultures are cheap and grow fast. Given this, subsequent cloning of the fusion fragments into the final target plasmids will be needed (e.g., expression vectors as pRK415 in bacteria or pCMV in mammalian cells).
CAPRRESI was tested for fusing two primary sigma factor genes: E. colirpoD and Rhizobium etlisigA. Primary sigma factors are RNA polymerase subunits responsible for transcription initiation, and they consist of four domains (i.e., &#963;1, &#963;2, &#963;3, and &#963;4)8. The amino acid sequence length of proteins encoded by rpoD and sigA are 613 and 685, respectively. RpoD and SigA share 48% identity (98% coverage). These primary sigma factors were split into two complementary fragments between regions &#963;2 and &#963;3. Two chimeric genes were assembled according to this design: chimera 01 (RpoD&#963;1-&#963;2 + SigA&#963;3-&#963;4) and chimera 02 (SigA&#963;1-&#963;2 + RpoD&#963;3-&#963;4). DNA fusion products were verified by sequencing.

<table><tbody><tr><td>Platinum Taq DNA polymerase High Fidelity</td><td>Thermo Fisher</td><td>11304011</td><td>Produces a mix of blunt/3&amp;rsquo;-A overhang ended PCR products</td></tr><tr><td>High pure plasmid isolation kit</td><td>Roche</td><td>11754785001</td><td>Used for all plasmid purification reactions</td></tr><tr><td>High pure PCR product purification kit</td><td>Roche</td><td>11732676001</td><td>Used for all PCR purification reactions from agarose gels</td></tr><tr><td>AflII restriction enzyme</td><td>New England Biolabs</td><td>R0520L</td><td>Recognizes sequence 5&amp;rsquo;-CTTAAG-3&amp;rsquo; and cuts at 37 &amp;deg;C</td></tr><tr><td>KpnI-HF restriction enzyme</td><td>New England Biolabs</td><td>R3142L</td><td>Recognizes sequence 5&amp;rsquo;-GGTACC-3&amp;rsquo; and cuts at 37 &amp;deg;C</td></tr><tr><td>SpeI-HF restriction enzyme</td><td>New England Biolabs</td><td>R3133L</td><td>Recognizes sequence 5&amp;rsquo;-ACTAGT-3&amp;rsquo; and cuts at 37 &amp;deg;C</td></tr><tr><td>XbaI restriction enzyme</td><td>New England Biolabs</td><td>R0145L</td><td>Recognizes sequence 5&amp;rsquo;-TCTAGA-3&amp;rsquo; and cuts at 37 &amp;deg;C</td></tr><tr><td>Ampicillin sodium salt</td><td>Sigma Aldrich</td><td>A0166-5G</td><td>Antibiotics</td></tr><tr><td>Nalidixic acid</td><td>Sigma Aldrich</td><td>N8878-5G</td><td>Antibiotics</td></tr><tr><td>Yeast Extract</td><td>Sigma Aldrich</td><td>Y1625-1KG</td><td>Bacterial cell culture</td></tr><tr><td>Casein peptone</td><td>Sigma Aldrich</td><td>70171-500G</td><td>Bacterial cell culture</td></tr><tr><td>NaCl</td><td>Sigma Aldrich</td><td>S9888-1KG</td><td>Sodium chloride</td></tr><tr><td>Agar</td><td>Sigma Aldrich</td><td>05040-1KG</td><td>Bacterial cell culture</td></tr><tr><td>&lt;em&gt;XbaI&lt;/em&gt; restriction enzyme</td><td>Thermo Fisher</td><td>FD0684</td><td>Fast digest &lt;em&gt;XbaI&lt;/em&gt; enzyme</td></tr><tr><td>&lt;em&gt;KpnI&lt;/em&gt; restriction enzyme</td><td>Thermo Fisher</td><td>FD0524</td><td>Fast digest &lt;em&gt;KpnI &lt;/em&gt;enzyme</td></tr><tr><td>Quick Ligation Kit</td><td>New England Biolabs</td><td>M2200S</td><td>Fast DNA ligation kit</td></tr><tr><td>&lt;em&gt;AflII&lt;/em&gt; restriction enzyme</td><td>Thermo Fisher</td><td>FD0834</td><td>Fast digest &lt;em&gt;AflII &lt;/em&gt;enzyme</td></tr><tr><td>&lt;em&gt;SpeI&lt;/em&gt; restriction enzyme</td><td>Thermo Fisher</td><td>FD1253</td><td>Fast digest &lt;em&gt;SpeI &lt;/em&gt;enzyme</td></tr></tbody></table>

caprresi: chimera assembly by plasmid recovery and restriction enzyme site insertion

Here, we present chimera assembly by plasmid recovery and restriction enzyme site insertion (CAPRRESI). CAPRRESI benefits from many strengths of the original plasmid recovery method and introduces restriction enzyme digestion to ease DNA ligation reactions (required for chimera assembly). For this protocol, users clone wildtype genes into the same plasmid (pUC18 or pUC19). After the in silico selection of amino acid sequence regions where chimeras should be assembled, users obtain all the synonym DNA sequences that encode them. Ad hoc Perl scripts enable users to determine all synonym DNA sequences. After this step, another Perl script searches for restriction enzyme sites on all synonym DNA sequences. This in silico analysis is also performed using the ampicillin resistance gene (ampR) found on pUC18/19 plasmids. Users design oligonucleotides inside synonym regions to disrupt wildtype and ampR genes by PCR. After obtaining and purifying complementary DNA fragments, restriction enzyme digestion is accomplished. Chimera assembly is achieved by ligating appropriate complementary DNA fragments. pUC18/19 vectors are selected for CAPRRESI because they offer technical advantages, such as small size (2,686 base pairs), high copy number, advantageous sequencing reaction features, and commercial availability. The usage of restriction enzymes for chimera assembly eliminates the need for DNA polymerases yielding blunt-ended products. CAPRRESI is a fast and low-cost method for fusing protein-coding genes.

Figure 1 depicts CAPRRESI. Using this method, two chimeric genes were assembled by exchanging the domains of two bacterial primary sigma factors (i.e., E. coli RpoD and R. etli SigA). The DNA sequences of the rpoD and sigA genes were obtained using the Artemis Genome Browser14 from GenBank genome files NC_000913 and NC_007761, respectively. The DNA sequence of the pUC18 vector was obtained from the nucleotide dat...

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CAPRRESI: Chimera Assembly by Plasmid Recovery and Restriction Enzyme Site Insertion

Here, we present chimera assembly by plasmid recovery and restriction enzyme site insertion (CAPRRESI), a protocol based on the insertion of restriction enzyme sites into synonym DNA sequences and functional plasmid recovery. This protocol is a fast and low-cost method for fusing protein-coding genes.

Here, we present chimera assembly by plasmid recovery and restriction enzyme site insertion (CAPRRESI). CAPRRESI benefits from many strengths of the ...

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11.5K Views.  Universidad Nacional Autónoma de México.  Hier presenteren we chimera assemblage door plasmideherstel en restrictie-enzymplaatsinzending (CAPRRESI), een protocol gebaseerd op de invoeging van restrictie-enzymplaatsen in synoniem-DNA-sequenties en functioneel plasmideherstel. Dit protocol is een snelle en goedkope methode voor het smelten van proteïne-coderende genen. 

Video: CAPRRESI: Chimera Assembly by Plasmid Recovery and Restriction Enzyme Site Insertion

Identification of Functional Protein Regions Through Chimeric Protein Construction

The goal of this protocol encompasses the design of chimeric proteins in which distinct regions of a protein are replaced by their corresponding sequences in a structurally similar protein, in order to determine the functional importance of these regions. Such chimeras are generated by means of a nested PCR protocol using overlapping DNA fragments and adequately designed primers, followed by their expression within a mammalian system to ensure native secondary structure and post-translational modifications. 
The functional role of a distinct region is then indicated by a loss of activity of the chimera in an appropriate readout assay. In consequence, regions harboring a set of critical amino acids are identified, which can be further screened by complementary techniques (e.g. site-directed mutagenesis) to increase molecular resolution. Although limited to cases in which a structurally related protein with differing functions can be found, chimeric proteins have been successfully employed to identify critical binding regions in proteins such as cytokines and cytokine receptors. This method is particularly suitable in cases in which the protein's functional regions are not well defined, and constitutes a valuable first step in directed evolution approaches to narrow down the regions of interest and reduce the screening effort involved.

Structurally related proteins frequently exert distinct biological functions. The exchange of equivalent regions of these proteins in order to create chimeric proteins constitutes an innovative approach to identify critical protein regions that are responsible for their functional divergence.

The goal of this protocol encompasses the design of chimeric proteins in which distinct regions of a protein are replaced by their corresponding sequences ...

Biochemistry

Homemade Site Directed Mutagenesis of Whole Plasmids

Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. With this method the cloned target gene can be altered by substitution, deletion or insertion of a few bases directly into a plasmid. It works by simply amplifying the whole plasmid, in a non PCR-based thermocycling reaction. During the reaction mutagenic primers, carrying the desired mutation, are integrated into the newly synthesized plasmid. In this video tutorial we demonstrate an easy and cost effective way to introduce base substitutions into a plasmid. The protocol works with standard reagents and is independent from commercial kits, which often are very expensive. Applying this protocol can reduce the total cost of a reaction to an eighth of what it costs using some of the commercial kits. In this video we also comment on critical steps during the process and give detailed instructions on how to design the mutagenic primers.

Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. Here we demonstrate an easy and cost effective way to introduce base substitutions into a plasmid using standard reagents.

Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. With this method the cloned ...

Implementing Gibson Assembly to Enhance Molecular Biology Education and Skills

Improving Student Outcomes with an Adaptable Molecular Cloning Course-Based Undergraduate Research Experience

The continuous advancement of molecular biology techniques requires that molecular biology curricula are&#160;regularly refined to effectively prepare students to enter the workforce with modern competencies. In particular, the emergence of Gibson Assembly, a highly customizable and adaptive molecular cloning technique, has advanced the landscape of molecular cloning in numerous research environments. Thus, we created a Gibson Assembly cloning module for deployment in a molecular biology laboratory course at California Polytechnic State University, San Luis Obispo and evaluated student learning outcomes from the module. Over three iterations of the course, students participated in an experiment-based independent project that involved cloning three unique plasmid libraries to support research projects in natural products biosynthesis. Students were given pre- and post-questionnaires to evaluate their understanding of molecular cloning and their confidence in molecular biology terms and techniques. Students&#8217; responses showed a significant increase in both learning molecular cloning concepts and in self-reported confidence with molecular cloning terms and techniques. This module framework can be generalized to teach Gibson Assembly for various applications, providing instructors with a toolkit for teaching an adaptable and emergent cloning technology while advancing their research projects.

An adaptable Gibson Assembly molecular cloning module was employed in a course-based undergraduate research experience (CURE) format for molecular biology laboratory course students. Assessment of student learning outcomes showed improved understanding and confidence in molecular cloning after completion of the CURE, and novel plasmids were cloned for natural product biosynthesis research.&#160;&#160;

The continuous advancement of molecular biology techniques requires that molecular biology curricula are&#160;regularly refined to effectively prepare ...

Rapid Assembly of Multi-Gene Constructs using Modular Golden Gate Cloning

The Golden Gate cloning method enables the rapid assembly of multiple genes in any user-defined arrangement. It utilizes type IIS restriction enzymes that cut outside of their recognition sites and create a short overhang. This modular cloning (MoClo) system uses a hierarchical workflow in which different DNA parts, such as promoters, coding sequences (CDS), and terminators, are first cloned into an entry vector. Multiple entry vectors then assemble into transcription units. Several transcription units then connect into a multi-gene plasmid. The Golden Gate cloning strategy is of tremendous advantage because it allows scar-less, directional, and modular assembly in a one-pot reaction. The hierarchical workflow typically enables the facile cloning of a large variety of multi-gene constructs with no need for sequencing beyond entry vectors. The use of fluorescent protein dropouts enables easy visual screening. This work provides a detailed, step-by-step protocol for assembling multi-gene plasmids using the yeast modular cloning (MoClo) kit. We show optimal and suboptimal results of multi-gene plasmid assembly and provide a guide for screening for colonies. This cloning strategy is highly applicable for yeast metabolic engineering and other situations in which multi-gene plasmid cloning is required.

The goal of this protocol is to provide a detailed, step-by-step guide for assembling multi-gene constructs using the modular cloning system based on Golden Gate cloning. It also gives recommendations on critical steps to ensure optimal assembly based on our experiences.

The Golden Gate cloning method enables the rapid assembly of multiple genes in any user-defined arrangement. It utilizes type IIS restriction enzymes that ...

Bioengineering

Markerless Gene Deletion by Floxed Cassette Allelic Exchange Mutagenesis in Chlamydia trachomatis

Chlamydia trachomatis is an obligate intracellular pathogen that has been historically difficult to genetically manipulate. Definitive progress in elucidating the mechanisms that C. trachomatis use to create and maintain a privileged intracellular niche has been limited due to a lack of genetic tools. Fortunately, there have recently been several new advances in genetic manipulation techniques. Among these is the development of fluorescence-reported allelic exchange mutagenesis (FRAEM). This method allows targeted gene deletion coupled with insertion of a selection cassette encoding antibiotic resistance and green fluorescent protein (GFP). Reliance on this strategy can be complicated when targeting genes within polycistronic operons due to the potential of polar effects on downstream genes. Floxed cassette allelic exchange mutagenesis (FLAEM), the protocol for which is described here, was developed to alleviate cassette-induced polar effects. FLAEM utilizes Cre-loxP genome editing to remove the selection cassette after targeted deletion by allelic exchange. The resulting strains contain markerless gene deletions of one or more coding sequences. This technique facilitates direct assessment of gene function and expands the repertoire of tools for genetic manipulation in C. trachomatis.

Markerless Gene Deletion by Floxed Cassette Allelic Exchange Mutagenesis in Chlamydia trachomatis

Described here is a method for targeted, markerless gene deletion in Chlamydia trachomatis using floxed cassette allelic exchange mutagenesis, FLAEM.

Chlamydia trachomatis is an obligate intracellular pathogen that has been historically difficult to genetically manipulate. Definitive progress in ...

Immunology and Infection

Construction of Adenoviral Vectors using DNA Assembly Technology

Adenoviral vectors have been used as a gene transfer tool in gene therapy for more than three decades. Here, we introduce a protocol to construct an adenoviral vector by manipulating the genomic DNA of wild-type HAdV-7 by using a DNA assembly method. First, an infectious clone of HAdV-7, pKan-Ad7, was generated by fusing the viral genomic DNA with a PCR product from plasmid backbone, comprising of the kanamycin-resistant gene and the origin of replication (Kan-Ori), through DNA assembly. This was done by designing a pair of PCR primers, that contained ~25 nucleotides of the terminal sequence of HAdV-7 inverted terminal repeat (ITR) at the 5' end, a non-cutter restriction enzyme site for HAdV-7 genome in the middle, and a template-specific sequence for PCR priming at the 3' end. Second, an intermediate plasmid-based strategy was employed to replace the E3 region with transgene-expressing elements in the infectious clone to generate an adenoviral vector. Briefly, pKan-Ad7 was digested with dual-cutter restriction enzyme Hpa I, and the fragment containing the E3 region was ligated to another PCR product of plasmid backbone by Gibson assembly to construct an intermediate plasmid pKan-Ad7HpaI. For convenience, restriction-assembly was used to designate the plasmid cloning method of combined restriction digestion and assembly. Using restriction-assembly, the E3 genes in pKan-Ad7HpaI was replaced with a green fluorescent protein (GFP) expression cassette, and the modified E3 region was released from the intermediate plasmid and restored to the infectious clone to generate an adenoviral plasmid pKAd7-E3GFP. Finally, pKAd7-E3GFP was linearized by Pme I digestion and used to transfect HEK293 packaging cells to rescue recombinant HAdV-7 virus. To conclude, a DNA assembly-based strategy was introduced here for constructing adenoviral vectors in general laboratories of molecular biology without the need of specialized materials and instruments.

In this study, an infectious clone of human adenovirus type 7 (HAdV-7) was constructed, and an E3-deleted HAdV-7 vector system was established by modifying the infectious clone. This strategy used here can be generalized to make gene transfer vectors from other wild-type adenoviruses.

Adenoviral vectors have been used as a gene transfer tool in gene therapy for more than three decades. Here, we introduce a protocol to construct an ...

Use of In Vivo Assembly for High-efficiency Plasmid Construction

In vivo assembly (IVA) is a molecular cloning method that uses intrinsic enzymes present in bacteria that promote intermolecular recombination of DNA fragments to assemble plasmids. This method functions by transforming DNA fragments with regions of 15-50 bp of homology into commonly used laboratory Escherichia coli strains and the bacteria use the RecA-independent repair pathway to assemble the DNA fragments into a plasmid. This method is more rapid and cost-effective than many molecular cloning methods that rely on in vitro assembly of plasmids prior to transformation into E. coli strains. This is because in vitro methods require the purchase of specialized enzymes and the performance of sequential enzymatic reactions that require incubations. However, unlike in vitro methods, IVA has not been experimentally shown to assemble linear plasmids. Here we share the IVA protocol used by our laboratory to rapidly assemble plasmids and subclone DNA fragments between plasmids with different origins of replication and antibiotic resistance markers.

In vivo assembly is a ligation-independent cloning method that relies on intrinsic DNA repair enzymes in bacteria to assemble DNA fragments by homologous recombination. This protocol is both time and cost-effective, as few reagents are required, and cloning efficiency can be as high as 99 %.

In vivo assembly (IVA) is a molecular cloning method that uses intrinsic enzymes present in bacteria that promote intermolecular recombination of DNA ...

Subcloning Plus Insertion (SPI) - A Novel Recombineering Method for the Rapid Construction of Gene Targeting Vectors

Gene targeting refers to the precise modification of a genetic locus using homologous recombination. The generation of novel cell lines and transgenic mouse models using this method necessitates the construction of a &#8216;targeting&#8217; vector, which contains homologous DNA sequences to the target gene, and has for many years been a limiting step in the process. Vector construction can be performed in vivo in Escherichia coli cells using homologous recombination mediated by phage recombinases using a technique termed recombineering. Recombineering is the preferred technique to subclone the long homology sequences (>4kb) and various targeting elements including selection markers that are required to mediate efficient allelic exchange between a targeting vector and its cognate genomic locus. Typical recombineering protocols follow an iterative scheme of step-wise integration of the targeting elements and require intermediate purification and transformation steps. Here, we present a novel recombineering methodology of vector assembly using a multiplex approach. Plasmid gap repair is performed by the simultaneous capture of genomic sequence from mouse Bacterial Artificial Chromosome libraries and the insertion of dual bacterial and mammalian selection markers. This subcloning plus insertion method is highly efficient and yields a majority of correct recombinants. We present data for the construction of different types of conditional gene knockout, or knock-in, vectors and BAC reporter vectors that have been constructed using this method. SPI vector construction greatly extends the repertoire of the recombineering toolbox and provides a simple, rapid and cost-effective method of constructing these highly complex vectors.

Subcloning Plus Insertion SPI - A Novel Recombineering Method for the Rapid Construction of Gene Targeting Vectors

Gene targeting methodologies can be used to generate transgenic mice with knockout, knock-in and tagged alleles. Here, we describe an improved method of recombineering in E. coli, that we term &#8216;subcloning plus insertion&#8217;, which can be used to generate custom gene targeting vectors rapidly.

Gene targeting refers to the precise modification of a genetic locus using homologous recombination. The generation of novel cell lines and transgenic ...

Generation of Plasmid Vectors Expressing FLAG-tagged Proteins Under the Regulation of Human Elongation Factor-1&#945; Promoter Using Gibson Assembly

Gibson assembly (GA) cloning offers a rapid, reliable, and flexible alternative to conventional DNA cloning methods. We used GA to create customized plasmids for expression of exogenous genes in mouse embryonic stem cells (mESCs). Expression of exogenous genes under the control of the SV40 or human cytomegalovirus promoters diminishes quickly after transfection into mESCs. A remedy for this diminished expression is to use the human elongation factor-1 alpha (hEF1&#945;) promoter to drive gene expression. Plasmid vectors containing hEF1&#945; are not as widely available as SV40- or CMV-containing plasmids, especially those also containing N-terminal 3xFLAG-tags. The protocol described here is a rapid method to create plasmids expressing FLAG-tagged CstF-64 and CstF-64 mutant under the expressional regulation of the hEF1&#945; promoter. GA uses a blend of DNA exonuclease, DNA polymerase and DNA ligase to make cloning of overlapping ends of DNA fragments possible. Based on the template DNAs we had available, we designed our constructs to be assembled into a single sequence. Our design used four DNA fragments: pcDNA 3.1 vector backbone, hEF1&#945; promoter part 1, hEF1&#945; promoter part 2 (which contained 3xFLAG-tag purchased as a double-stranded synthetic DNA fragment), and either CstF-64 or specific CstF-64 mutant. The sequences of these fragments were uploaded to a primer generation tool to design appropriate PCR primers for generating the DNA fragments. After PCR, DNA fragments were mixed with the vector containing the selective marker and the GA cloning reaction was assembled. Plasmids from individual transformed bacterial colonies were isolated. Initial screen of the plasmids was done by restriction digestion, followed by sequencing. In conclusion, GA allowed us to create customized plasmids for gene expression in 5 days, including construct screens and verification.

Synthesis of custom plasmids is labor and time consuming. This protocol describes the use of Gibson assembly cloning to reduce the work and duration of custom DNA cloning procedure. The protocol described also produces reliable tagged protein constructs for mammalian expression at similar cost to the traditional cut-and-paste DNA cloning.

Gibson assembly (GA) cloning offers a rapid, reliable, and flexible alternative to conventional DNA cloning methods. We used GA to create customized ...

Generation of Marked and Markerless Mutants in Model Cyanobacterial Species

Cyanobacteria are ecologically important organisms and potential platforms for production of biofuels and useful industrial products. Genetic manipulation of cyanobacteria, especially model organisms such as Synechocystis sp. PCC6803 and Synechococcus sp. PCC7002, is a key tool for both basic and applied research. Generation of unmarked mutants, whereby chromosomal alterations are introduced into a strain via insertion of an antibiotic resistance cassette (a manipulatable fragment of DNA containing one or more genes), followed by subsequent removal of this cassette using a negative selectable marker, is a particularly powerful technique. Unmarked mutants can be repeatedly genetically manipulated, allowing as many alterations to be introduced into a strain as desired. In addition, the absence of genes encoding antibiotic resistance proteins in the mutated strain is desirable, as it avoids the possibility of 'escape' of antibiotic resistant organisms into the environment. However, detailed methods for repeated rounds of genetic manipulation of cyanobacteria are not well described in the scientific literature. Here we provide a comprehensive description of this technique, which we have successfully used to generate mutants with multiple deletions, single point mutations within a gene of interest and insertion of novel gene cassettes.

Introducing multiple genomic alterations into cyanobacteria is an essential tool in the development of strains for industrial and basic research purposes. We describe a system for generating unmarked mutants in the model cyanobacterial species Synechocystis sp. PCC6803 and marked mutants in Synechococcus sp. PCC7002.

Cyanobacteria are ecologically important organisms and potential platforms for production of biofuels and useful industrial products. Genetic manipulation ...

CAPRRESI: Chimera Assembly by Plasmid Recovery and Restriction Enzyme Site Insertion

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