We greatly appreciate the valuable advice provided by Prof. J.P. van Pijkeren (University of Wisconsin-Madison), whose guidance on working with L. reuteri ATCC PTA 6475 provided a foundation for the methods described here.

1. Preparing L. reuteri DSM20016 electrocompetent cells
NOTE: This is based on a protocol by Berthier et al.17, with centrifugation speeds informed by Rattanachaikunsopon et al.18.
<ol>
	<li>In a 50 mL centrifuge tube, inoculate L. reuteri from glycerol stock into 6 mL of deMan Rogosa Sharpe (MRS) broth. Incubate aerobically overnight at 37 &#176;C in a static incubator.</li>
	<li>The next morning, inoculate 4 mL of the overnight culture into 200 mL of MRS broth (1:50 dilution).</li>
	<li>Allow to grow aerobically in a static 37 &#176;C incubator until the 600 nm optical density value (OD600) reaches 0.5-0.85; this should take roughly 2-3 h.</li>
	<li>As soon as an adequate OD600 value is reached, decant the media into 50 mL centrifuge tubes and place on ice while balancing the tubes.</li>
	<li>Centrifuge for 5 min at 5,000 x g in a pre-chilled, 4 &#176;C centrifuge.</li>
	<li>Discard the supernatant, resuspend each pellet in 50 mL of pre-chilled (0 &#176;C to 4 &#176;C) ddH2O, and centrifuge again with the same settings as the previous step. Keep the cells on ice as much as possible.</li>
	<li>Repeat step 1.6.</li>
	<li>Resuspend each pellet in 25 mL of ddH2O: 0.5 M sucrose, 10% glycerol. Centrifuge at 5,000 x g at 4 &#176;C for 10 min. Discard the supernatant and put the cells back on ice.</li>
	<li>Resuspend all the pellets in the same 2 mL of ddH2O: 0.5 M sucrose, 10% glycerol.</li>
	<li>Aliquot into 50 &#181;L to 100 &#181;L portions in pre-chilled microcentrifuge tubes; store at -80 &#176;C for later use.</li>
</ol>
2. Electroporation of L. reuteri 
NOTE: Avoid pipetting as much as possible in the following steps. Inclusion of a control electroporation, with no plasmid added, is advised to ensure the antibiotic selection is adequate.
<ol>
	<li>Take an entire electrocompetent L. reuteri aliquot and thaw it on ice.</li>
	<li>Gently mix 5 &#181;L to 10 &#181;L of plasmid (final plasmid concentration &#62;6 nM) into the thawed aliquot, avoiding pipetting as much as possible.</li>
	<li>Transfer to an ice-chilled, 1 mm gap electroporation cuvette.</li>
	<li>Electroporate at 1.25 kV, 400 &#8486;, and 25 &#181;F.</li>
	<li>Add 1 mL of room temperature (RT) MRS broth and mix by inverting the cuvette once or twice.</li>
	<li>Place the cuvettes into a static incubator at 37 &#176;C for 2.5-3 h to allow for recovery.</li>
	<li>Plate the entire amount onto multiple MRS agar plates with appropriate selection.</li>
	<li>Place the plates inside a completely airtight container with a small lit candle (&#34;tealight&#34;) and an anaerobic atmosphere-generating sachet.</li>
	<li>Grow at 37 &#176;C for 2-3 days or until visible colonies are present.</li>
</ol>
3. Measurement of the acid-resistant fluorescent reporter protein mCherry2
<ol>
	<li>Pick any colonies required for measurement and inoculate into a 96-well storage microplate with 1.5 mL of filter sterilized (non-autoclaved) MRS broth per well and an appropriate selection antibiotic.</li>
	<li>Incubate aerobically for 24 h overnight at 37 &#176;C without shaking. 
	NOTE: This storage microplate should be kept for use in section 4 (colony PCR).</li>
	<li>L. reuteri will precipitate out of the media when in the stationary phase; resuspend the overnight culture via pipetting.</li>
	<li>Transfer 200 &#181;L into a flat, clear-bottom, 96-well plate, transfer the plate to a plate reader, and measure the OD and fluorescence of mCherry2 (excitation [Ex]: 589 nm; emission [Em]: 610 nm) or other relevant reporters.</li>
</ol>
4. Confirmation of plasmid uptake via colony PCR
<ol>
	<li>From the 96-well storage microplate from section 3, transfer 5 &#181;L to a PCR tube. 
	NOTE: If &#62;5 min have passed, it may be necessary to resuspend the L. reuteri a second time.</li>
	<li>In a portable benchtop microcentrifuge, spin down until the pellet can be seen (roughly 2 min at 2,000 x g), discard the supernatant, and resuspend the pellet in 20 &#181;L of 20 mM NaOH.</li>
	<li>Boil at 95 &#176;C for 5 min, vortex, and repeat the boil a second time.</li>
	<li>Immediately chill the samples; try to keep the samples on ice as much as possible in the following steps to lower the likelihood of template degradation and PCR inhibition.</li>
	<li>Spin down in a portable benchtop microcentrifuge at 2,000 x g for 2 min until the cell debris is pelleted, then take 1 &#181;L of the supernatant and dilute into 99 &#181;L of ice-cold DNase- and RNase-free ddH2O (100x dilution).</li>
	<li>Use the 100x dilution as the template DNA in a standard PCR reaction using plasmid-specific primers. Include a positive control with a plasmid derived from the E. coli mini-prep.</li>
	<li>Return the samples on ice and add an appropriate loading dye at a 1x concentration.</li>
	<li>Run in 1% agarose gel in TAE buffer (tris-acetate-ethylenediaminetetraacetic acid [EDTA]) at 110 V for 30 min. Image if necessary.</li>
</ol>
5. Mini-prep protocol for L. reuteri , followed by PCR to confirm plasmid presence
NOTE: Protocol intended for use with the mini-prep kit listed in the Table of Materials.
<ol>
	<li>Inoculate L. reuteri into 10 mL of MRS broth containing an appropriate antibiotic and incubate overnight aerobically at 37 &#176;C in a static incubator.</li>
	<li>Centrifuge at 5,000 x g for 10 min in a pre-chilled 4 &#176;C centrifuge.</li>
	<li>Wash the pellet in 2 mL of standard P1 buffer (included with the kit)&#160;in order to negate acidity that may interfere with later steps, centrifuge with the same setting as previously described, and discard the supernatant.</li>
	<li>Resuspend the pellet in 250 &#181;L of modified P1 buffer containing 10 mg/mL lysozyme and 100 U/mL mutanolysin in order to lyse bacterial cells. Incubate for 1-2 h at 37 &#176;C.</li>
	<li>Add 250 &#181;L of buffer P2, mix by inverting four to six times, and incubate at RT for no longer than 5 min.</li>
	<li>Add 350 &#181;L of buffer N3 (included with the kit) and immediately invert four to six times gently to mix.</li>
	<li>Centrifuge at 10,000 x g for 10 min.</li>
	<li>Transfer as much supernatant as possible to a spin column and centrifuge at 10,000 x g for 60 s. Discard the flow through.</li>
	<li>Wash the spin column with 500 &#181;L of buffer PB (included with the kit) and centrifuge at 10,000 x g for 60 s. Discard the flow through.</li>
	<li>Wash the spin column with 750 &#181;L of buffer PE (included with the kit), and centrifuge at 10,000 x g for 60 s. Discard the flow through and centrifuge for 60 s to remove any residual buffer.</li>
	<li>Place the spin column in a 1.5 mL microcentrifuge tube. Apply 20-30 &#181;L of DNAse- and RNAse-free ddH2O to the filter of the spin column and leave for 1-2 min, before centrifuging at 10,000 x g for 60 s.</li>
	<li>Perform a standard PCR reaction using plasmid-specific primers (pTRKH3_pTUSeq_F: CACCCGTTCTCGGAGCA, pTRKH3_pTUSeq_R: CTACGAGTTGCATGATAAAGAAGACA), with eluate providing the template DNA. Include a positive control with a plasmid derived from the E. coli mini-prep. 
	NOTE: The PCR settings used in this study are: 98 &#176;C for 5 min, (98 &#176;C for 30 s, 55 &#176;C for 30 s, 72 &#176;C for 45 s) 30 cycles, 72 &#176;C for 2 min, 4 &#176;C hold. The parameters are highly dependent on the polymerase used, fragment length, and exact primers used by any person utilizing this protocol.</li>
</ol>

Measurement of Reporter Proteins in Limosilactobacillus reuteri DSM20016

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No conflicts of interest exist.

The most critical step for the transformation of L. reuteri DSM20016 is the generation of anaerobic growth conditions after transformations are plated; colonies gained in aerobic conditions are only very occasional and generally fail to grow when inoculated in MRS broth. Plating the entire recovery volume should also be practiced to maximize the probability of colony growth. Even with these two critical steps, transformation efficiency is still a limitation on experimentation, as expected colonies can number as ...

The genus Lactobacillus were historically classified as gram-positive, rod-shaped, non-spore-forming, either facultative anaerobes or microaerophiles that break sugars down to primarily produce lactic acid1. These loose criteria led to Lactobacillus being, phenotypically and genotypically, an extremely diverse genus. This broad categorization resulted in the genus being reclassified, introducing 23 novel genera in 20202.
The old, broader genus included major commensal and probiotic species generally regarded as safe (GRAS) for consumption3. The Lactobacillaceae family maintains a public perception of being &#39;good bacteria&#39; due to many reported health benefits bestowed via the consumption of various strains4,5,6,7. The ease with which they can navigate the gastrointestinal tract8 and their public acceptance combine to position Lactobacillaceae strains as strong candidates as chassis organisms for ingestible medicinal, therapeutic, or diagnostic applications.
The wide range of characteristics present within the Lactobacillaceae family has led to a situation in which there is no de facto model-organism strain; research groups have tended to select species with the properties most relevant to their particular aims. (For example, dairy fermentation labs could choose L. lactis; studies of vegetable fermentation might select L. plantarum; research on probiotics might focus on L. acidophilus; and so on.)
This same wide range of characteristics across species has led to an accumulation of protocols and procedures that may work well for one subset of the Lactobacillaceae family, but require optimization to work efficiently (or perhaps to function at all) in others9. This need for optimization between family members and even within members of the same species can frustrate the efforts of unfamiliar researchers. Protocols published in the methods sections of papers can also include their own modifications10, leading to fragmented, decentralized protocol collections.
L. reuteri is considered a widely vertebrate commensal, found consistently in mammalian, avian11 and fish12 gastrointestinal (GI) tracts. L. reuteri sub-strains are often genetically specialized, via mucus adhesion protein adaptation, to more permanently colonize specific native hosts8,11,13. GI tract Limosilactobacillus species can be isolated in hosts outside their native host, but tend more toward a transient nature8.
Due to human-host specialization, L. reuteri DSM20016 positions itself very well as a chassis for diagnostic or therapeutic applications at any point in the human GI tract, and the strain DSM20016 could provide a longer-lasting window of effect for interventions when compared to more transitory strains.
In this paper, we outline a series of protocols with demonstrated effectiveness in Limosilactobacillus reuteri (strain designation: F275; other collection numbers: DSM20016, ATCC23272, CIP109823), along with centralized information on the strain from other sources to aid in molecular and systems biology applications. Procedures laid out herein should enable a researcher with no prior experience to culture L. reuteri, create electrocompetent stocks, select transformed colonies, confirm transformation via colony polymerase chain reaction (PCR), and measure designed system response via fluorescent reporter proteins.
We note that related protocols have covered CRISPR-Cas9 assisted ssDNA genome recombineering in L. reuteri (strain: ATCC-PTA-6475)14, and CRSIPR-Cas9 nickase-assisted genome editing in multiple non-L. reuteri, Lactobacillaceae family stains15,16; these do not, however, address the L. reuteri DSM20016 strain that is our focus here.

<table><tbody><tr><td>1 kb Plus DNA Ladder</td><td>NEB</td><td>N3200L</td><td></td></tr><tr><td>1mL Spectrophotometer cuvettes</td><td>Thomas Scientific</td><td>1145J12</td><td></td></tr><tr><td>Agarose&amp;nbsp;</td><td>BioShop</td><td>AGR001</td><td></td></tr><tr><td>Allegra X-15R (refrigerated centrifuge)</td><td>Beckman Allegra&amp;nbsp;</td><td>N/A</td><td>No longer in production</td></tr><tr><td>AnaeroGen 2.5 L Sachet</td><td>Thermo Scientific</td><td>OXAN0025A</td><td></td></tr><tr><td>BTX, ECM 399 electroporation system</td><td>VWR</td><td>58017-984</td><td></td></tr><tr><td>Centrifuge tubes (50 mL)</td><td>FroggaBio</td><td>TB50-500</td><td></td></tr><tr><td>DNA gel x6 loading dye</td><td>NEB</td><td>B7024S</td><td></td></tr><tr><td>Electroporation cuvette</td><td>Fisherbrand</td><td>FB101</td><td></td></tr><tr><td>Erythromycin</td><td>Millipore Sigma</td><td>E5389-5G</td><td></td></tr><tr><td>Gel electroporation bath/dock</td><td>VWR</td><td>76314-748</td><td></td></tr><tr><td>Glycerol&amp;nbsp;</td><td>BioShop</td><td>GLY001</td><td></td></tr><tr><td>Limosilactobacillus reuteri</td><td>Leibniz Institute DSMZ</td><td>DSM20016</td><td>Strain designation F275</td></tr><tr><td>Lysozyme</td><td>BioShop</td><td>LYS702.5</td><td></td></tr><tr><td>Microcentrifuge tubes (1.7 mL)</td><td>FroggaBio</td><td>LMCT1.7B</td><td></td></tr><tr><td>Miniprep kit (Qiagen)</td><td>Qiagen</td><td>27106</td><td>slpGFP replaced with constitutive, codon optimised, mCherry2 reporter protein&amp;nbsp;</td></tr><tr><td>MRS Broth (Dehydrated)</td><td>Thermo Scientific</td><td>CM0359B</td><td></td></tr><tr><td>Mutanolysin</td><td>Millipore Sigma</td><td>M9901-5KU</td><td></td></tr><tr><td>NaOH&amp;nbsp;</td><td>Millipore Sigma</td><td>1064691000</td><td></td></tr><tr><td>P100 Pipette</td><td>Eppendorf</td><td>3123000047</td><td></td></tr><tr><td>P1000 Pipette</td><td>Eppendorf</td><td>3123000063</td><td></td></tr><tr><td>P2.5 Pipette</td><td>Eppendorf</td><td>3123000012</td><td></td></tr><tr><td>P20 Pipette</td><td>Eppendorf</td><td>3123000039</td><td></td></tr><tr><td>P200 Pipette</td><td>Eppendorf</td><td>3123000055</td><td></td></tr><tr><td>PCR tubes</td><td>FroggaBio</td><td>STF-A120S</td><td></td></tr><tr><td>Personal benchtop microcentrifuge</td><td>Genlantis</td><td>E200100</td><td></td></tr><tr><td>Petri dishes</td><td>VWR</td><td>25384-088</td><td></td></tr><tr><td>PTC-150 Thermal Cycler</td><td>MJ Research</td><td>N/A</td><td>No longer in production</td></tr><tr><td>pTRKH3_slpGFP (modified)</td><td>Addgene</td><td>27168</td><td></td></tr><tr><td>SPECTRONIC 200 Spectrophotometer</td><td>Thermo Scientific</td><td>840-281700</td><td></td></tr><tr><td>Storage microplate</td><td>Fisher Scientific</td><td>14-222-225</td><td></td></tr><tr><td>Sucrose</td><td>BioShop</td><td>SUC507</td><td></td></tr><tr><td>TAE Buffer 50x</td><td>Thermo Scientific</td><td>B49</td><td></td></tr><tr><td>Vortex</td><td>VWR</td><td>58816-121</td><td>No longer in production</td></tr><tr><td>VWR 1500E incubator</td><td>VWR</td><td>N/A</td><td>No longer in production</td></tr></tbody></table>

methods for electroporation and transformation confirmation in limosilactobacillus reuteri dsm20016

Lactobacillus were an incredibly large, diverse genus of bacteria comprising 261 species, several of which were commensal strains with the potential for use as a chassis for synthetic biological endeavors within the gastrointestinal tract. The wide phenotypic and genotypic variation observed within the genus led to a recent reclassification and the introduction of 23 novel genera.
Due to the breadth of variations within the old genera, protocols demonstrated in one member may not work as advertised with other members. A lack of centralized information on how exactly to manipulate specific strains has led to a range of ad hoc approaches, often adapted from other bacterial families. This can complicate matters for researchers starting in the field, who may not know which information does or does not apply to their chosen strain.
In this paper, we aim to centralize a set of protocols with demonstrated success, specifically in&#160;the Limosilactobacillus reuteri strain designation F275 (other collection numbers: DSM20016, ATCC23272, CIP109823), along with troubleshooting advice and common issues one may encounter. These protocols should enable a researcher with little to no experience working with L. reuteri DSM20016 to transform a plasmid, confirm transformation, and measure system feedback in a plate reader via a reporter protein.

Transformation efficiencies 
L. reuteri does not require a dcm-/dam- non-methylated plasmid, as observed for other Lactobacillaceae19,20 (see Figure 1). Electroporation of L. reuteri DSM20016 with 10 &#181;L of the 8.5 kb plasmid pTRKH3_mCherry2 (pAM&#946;1 theta origin of replication) should give transformation efficiencies of roughly 80 colony forming units (CFU...

Watch this Scientific Journal Video about Methods for Electroporation and Transformation Confirmation in Limosilactobacillus reuteri DSM20016 at JoVE.com

Author Spotlight: Methods for Electroporation and Transformation Confirmation in Limosilactobacillus reuteri DSM20016  

Here, we present protocols for working with Limosilactobacillus reuteri DSM20016, detailing growth, plasmid transformation, colony PCR, fluorescent reporter protein measurement, and limited plasmid mini-prep, as well as common issues and troubleshooting. These protocols allow the measurement of reporter proteins in DSM20016, or confirmation via colony PCR if no reporter is involved.

Methods for Electroporation and Transformation Confirmation in Limosilactobacillus reuteri DSM20016

Lactobacillus were an incredibly large, diverse genus of bacteria comprising 261 species, several of which were commensal strains with the potential for ...

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3.5K Views.  University of Toronto Mississauga. Here, we present protocols for working with Limosilactobacillus reuteri DSM20016, detailing growth, plasmid transformation, colony PCR, fluorescent reporter protein measurement, and limited plasmid mini-prep, as well as common issues and troubleshooting. These protocols allow the measurement of reporter proteins in DSM20016, or confirmation via colony PCR if no reporter is involved. 

Video: Author Spotlight: Methods for Electroporation and Transformation Confirmation in Limosilactobacillus reuteri DSM20016  

Electroporation of Mycobacteria

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly caused by the thick and waxy cell wall, but is compounded by the fact that most molecular techniques have been developed for distantly-related species such as Escherichia coli and Bacillus subtilis. In spite of these obstacles, mycobacterial plasmids have been identified and DNA transformation of many mycobacterial species have now been described. The most successful method for introducing DNA into mycobacteria is electroporation. Many parameters contribute to successful transformation; these include the species/strain, the nature of the transforming DNA, the selectable marker used, the growth medium, and the  conditions for the electroporation pulse. Optimized methods for the transformation of both slow- and fast-grower are detailed here. Transformation efficiencies for different mycobacterial species and with various selectable markers are reported.

Mycobacterial pathogenic strategies remain poorly understood. The slow growth rate of most species, the impenetrable nature of the cell-wall, and the hazards of working with pathogens make mycobacteria difficult to study and are largely responsible for our poor understanding of these organisms. In this video we will demonstrate the technique of electroporation, which involves subjecting cells to a brief high electrical impulse to allow the entry of DNA. It is the most widely used method for introducing DNA into mycobacterial cells.

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly ...

Biology

Transformation of Probiotic Yeast and Their Recovery from Gastrointestinal Immune Tissues Following Oral Gavage in Mice

Development of recombinant oral therapy would allow for more direct targeting of the mucosal immune system and improve the ability to combat gastrointestinal disorders. Adapting probiotic yeast in particular for this approach carries several advantages. These strains have not only the potential to synthesize a wide variety of complex heterologous proteins but are also capable of surviving and protecting those proteins during transit through the intestine. Critically, however, this approach requires expertise in many diverse laboratory techniques not typically used in tandem. Furthermore, although individual protocols for yeast transformation are well characterized for commonly used laboratory strains, emphasis is placed here on alternative approaches and the importance of optimizing transformation for less well characterized probiotic strains. Detailing these methods will help facilitate discussion as to the best approaches for testing probiotic yeast as oral drug delivery vehicles and indeed serve to advance the development of this novel strategy for gastrointestinal therapy.

Presented here is a unified description of techniques that can be used to develop, transform, administer, and test heterologous protein expression of the probiotic yeast Saccharomyces boulardii.

Development of recombinant oral therapy would allow for more direct targeting of the mucosal immune system and improve the ability to combat ...

Immunology and Infection

Mapping Bacterial Functional Networks and Pathways in Escherichia Coli using Synthetic Genetic Arrays

Phenotypes are determined by a complex series of physical (e.g. protein-protein) and functional (e.g. gene-gene or genetic) interactions (GI)1. While physical interactions can indicate which bacterial proteins are associated as complexes, they do not necessarily reveal pathway-level functional relationships1. GI screens, in which the growth of double mutants bearing two deleted or inactivated genes is measured and compared to the corresponding single mutants, can illuminate epistatic dependencies between loci and hence provide a means to query and discover novel functional relationships2. Large-scale GI maps have been reported for eukaryotic organisms like yeast3-7, but GI information remains sparse for prokaryotes8, which hinders the functional annotation of bacterial genomes. To this end, we and others have developed high-throughput quantitative bacterial GI screening methods9, 10.
Here, we present the key steps required to perform quantitative E. coli Synthetic Genetic Array (eSGA) screening procedure on a genome-scale9, using natural bacterial conjugation and homologous recombination to systemically generate and measure the fitness of large numbers of double mutants in a colony array format. Briefly, a robot is used to transfer, through conjugation, chloramphenicol (Cm) - marked mutant alleles from engineered Hfr (High frequency of recombination) 'donor strains' into an ordered array of kanamycin (Kan) - marked F- recipient strains. Typically, we use loss-of-function single mutants bearing non-essential gene deletions (e.g. the 'Keio' collection11) and essential gene hypomorphic mutations (i.e. alleles conferring reduced protein expression, stability, or activity9, 12, 13) to query the functional associations of non-essential and essential genes, respectively. After conjugation and ensuing genetic exchange mediated by homologous recombination, the resulting double mutants are selected on solid medium containing both antibiotics. After outgrowth, the plates are digitally imaged and colony sizes are quantitatively scored using an in-house automated image processing system14. GIs are revealed when the growth rate of a double mutant is either significantly better or worse than expected9. Aggravating (or negative) GIs often result between loss-of-function mutations in pairs of genes from compensatory pathways that impinge on the same essential process2. Here, the loss of a single gene is buffered, such that either single mutant is viable. However, the loss of both pathways is deleterious and results in synthetic lethality or sickness (i.e. slow growth). Conversely, alleviating (or positive) interactions can occur between genes in the same pathway or protein complex2 as the deletion of either gene alone is often sufficient to perturb the normal function of the pathway or complex such that additional perturbations do not reduce activity, and hence growth, further. Overall, systematically identifying and analyzing GI networks can provide unbiased, global maps of the functional relationships between large numbers of genes, from which pathway-level information missed by other approaches can be inferred9.

Mapping Bacterial Functional Networks and Pathways in Escherichia Coli using Synthetic Genetic Arrays

Systematic, large-scale synthetic genetic (gene-gene or epistasis) interaction screens can be used to explore genetic redundancy and pathway cross-talk. Here, we describe a high-throughput quantitative synthetic genetic array screening technology, termed eSGA that we developed for elucidating epistatic relationships and exploring genetic interaction networks in Escherichia coli.

Phenotypes  are determined by a complex series of physical (e.g. protein-protein) and  functional (e.g. gene-gene or genetic) interactions (GI)1. While ...

Genomic Transformation of the Picoeukaryote Ostreococcus tauri

Common problems hindering rapid progress in Plant Sciences include cellular, tissue and whole organism complexity, and notably the high level of genomic redundancy affecting simple genetics in higher plants. The novel model organism Ostreococcus tauri is the smallest free-living eukaryote known to date, and possesses a greatly reduced genome size and cellular complexity1,2, manifested by the presence of just one of most organelles (mitochondrion, chloroplast, golgi stack) per cell, and a genome containing only ~8000 genes. Furthermore, the combination of unicellularity and easy culture provides a platform amenable to chemical biology approaches. Recently, Ostreococcus has been successfully employed to study basic mechanisms underlying circadian timekeeping3-6. Results from this model organism have impacted not only plant science, but also mammalian biology7. This example highlights how rapid experimentation in a simple eukaryote from the green lineage can accelerate research in more complex organisms by generating testable hypotheses using methods technically feasible only in this background of reduced complexity. Knowledge of a genome and the possibility to modify genes are essential tools in any model species. Genomic1, Transcriptomic8, and Proteomic9 information for this species is freely available, whereas the previously reported methods6,10 to genetically transform Ostreococcus are known to few laboratories worldwide.
In this article, the experimental methods to genetically transform this novel model organism with an overexpression construct by means of electroporation are outlined in detail, as well as the method of inclusion of transformed cells in low percentage agarose to allow selection of transformed lines originating from a single transformed cell. Following the successful application of Ostreococcus to circadian research, growing interest in Ostreococcus can be expected from diverse research areas within and outside plant sciences, including biotechnological areas. Researchers from a broad range of biological and medical sciences that work on conserved biochemical pathways may consider pursuing research in Ostreococcus, free from the genomic and organismal complexity of larger model species.

Genomic Transformation of the Picoeukaryote Ostreococcus tauri

This article describes genetic transformation of the unicellular marine alga Ostreococcus tauri by electroporation. This eukaryotic organism is an effective model platform for higher plants, possesing greatly reduced genomic and cellular complexity and being readily amenable to both cell culture and chemical biology.

Common problems hindering rapid progress in Plant Sciences include cellular, tissue and whole organism complexity, and notably the high level of genomic ...

Rapid Protocol for Preparation of Electrocompetent Escherichia coli and Vibrio cholerae

Electroporation has become a widely used method for rapidly and efficiently introducing foreign DNA into a wide range of cells. Electrotransformation has become the method of choice for introducing DNA into prokaryotes that are not naturally competent. Electroporation is a rapid, efficient, and streamlined transformation method that, in addition to purified DNA and competent bacteria, requires commercially available gene pulse controller and cuvettes. In contrast to the pulsing step, preparation of electrocompetent cells is time consuming and labor intensive involving repeated rounds of centrifugation and washes in decreasing volumes of sterile, cold water, or non-ionic buffers of large volumes of cultures grown to mid-logarithmic phase of growth. Time and effort can be saved by purchasing electrocompetent cells from commercial sources, but the selection is limited to commonly employed E. coli laboratory strains. We are hereby disseminating a rapid and efficient method for preparing electrocompetent E. coli, which has been in use by bacteriology laboratories for some time, can be adapted to V. cholerae and other prokaryotes. While we cannot ascertain whom to credit for developing the original technique, we are hereby making it available to the scientific community.

Rapid Protocol for Preparation of Electrocompetent Escherichia coli and Vibrio cholerae

Electroporation is a commonly employed method for introducing DNA into bacteria in a process known as transformation. Traditional protocols for the preparation of electrocompetent cells are time consuming and labor intensive. This article describes an alternate, rapid, and efficient method for the preparation of electrocompetent cells presently employed by some laboratories.

Electroporation has become a widely used method for rapidly and efficiently introducing foreign DNA into a wide range of cells. Electrotransformation has ...

Generation of Null Mutants to Elucidate the Role of Bacterial Glycosyltransferases in Bacterial Motility

The study of glycosylation in prokaryotes is a rapidly growing area. Bacteria harbor different glycosylated structures on their surface whose glycans constitute a strain-specific barcode. The associated glycans show higher diversity in sugar composition and structure than those of eukaryotes and are important in bacterial-host recognition processes and interaction with the environment. In pathogenic bacteria, glycoproteins have been involved in different stages of the infectious process, and glycan modifications can interfere with specific functions of glycoproteins. However, despite the advances made in the understanding of glycan composition, structure, and biosynthesis pathways, understanding of the role of glycoproteins in pathogenicity or interaction with the environment remains very limited. Furthermore, in some bacteria, the enzymes required for protein glycosylation are shared with other polysaccharide biosynthetic pathways, such as lipopolysaccharide and capsule biosynthetic pathways. The functional importance of glycosylation has been elucidated in several bacteria through mutation of specific genes thought to be involved in the glycosylation process and the study of its impact on the expression of the target glycoprotein and the modifying glycan. Mesophilic Aeromonas have a single and O-glycosylated polar flagellum. Flagellar glycans show diversity in carbohydrate composition and chain length between Aeromonas strains. However, all strains analyzed to date show a pseudaminic acid derivative as the linking sugar that modifies serine or threonine residues. The pseudaminic acid derivative is required for polar flagella assembly, and its loss has an impact on adhesion, biofilm formation, and colonization. The protocol detailed in this article describes how the construction of null mutants can be used to understand the involvement of genes or genome regions containing putative glycosyltransferases in the biosynthesis of a flagellar glycan. This includes the potential to understand the function of the glycosyltransferases involved and the role of the glycan. This will be achieved by comparing the glycan deficient mutant to the wild-type strain.

Here, we describe the construction of null mutants of Aeromonas in specific glycosyltransferases or regions containing glycosyltransferases, the motility assays, and flagella purification performed to establish the involvement and function of their encoded enzymes in the biosynthesis of a glycan, as well as the role of this glycan in bacterial pathogenesis.

The study of glycosylation in prokaryotes is a rapidly growing area. Bacteria harbor different glycosylated structures on their surface whose glycans ...

Single&#45;Copy Gene Expression for Characterizing Multidrug Efflux Systems

Characterizing Multidrug Efflux Systems in Acinetobacter baumannii Using an Efflux&#45;Deficient Bacterial Strain and a Single&#45;Copy Gene Expression System

Acinetobacter baumannii is recognized as a challenging Gram-negative pathogen due to its widespread resistance to antibiotics. It is crucial to comprehend the mechanisms behind this resistance to design new and effective therapeutic options. Unfortunately, our ability to investigate these mechanisms in A. baumannii is hindered by the paucity of suitable genetic manipulation tools. Here, we describe methods for utilizing a chromosomal mini-Tn7-based system to achieve single-copy gene expression in an A. baumannii strain that lacks functional RND-type efflux mechanisms. Single-copy insertion and inducible efflux pump expression are quite advantageous, as the presence of RND efflux operons on high-copy number plasmids is often poorly tolerated by bacterial cells. Moreover, incorporating recombinant mini-Tn7 expression vectors into the chromosome of a surrogate A. baumannii host with increased efflux sensitivity helps circumvent interference from other efflux pumps. This system is valuable not only for investigating uncharacterized bacterial efflux pumps but also for assessing the effectiveness of potential inhibitors targeting these pumps.

Author Spotlight: Advancing Antibiotic Resistance Research Using an Efflux&#45;Deficient Bacterial Strain and a Single&#45;Copy Gene Expression System

We describe a facile procedure for the single-copy chromosomal complementation of an efflux pump gene using a mini-Tn7-based expression system into an engineered efflux-deficient strain of Acinetobacter baumannii. This precise genetic tool allows for controlled gene expression, which is key for the characterization of efflux pumps in multidrug resistant pathogens.

Acinetobacter baumannii is recognized as a challenging Gram-negative pathogen due to its widespread resistance to antibiotics. It is crucial to comprehend ...

Bacterial Transformation: The Heat Shock Method

Transformation is the process that occurs when a cell ingests foreign DNA from its surroundings. Transformation can occur in nature in certain types of bacteria. In molecular biology, transformation is artificially reproduced in the lab via the creation of pores in bacterial cell membranes. Bacterial cells that are able to take up DNA from the environment are called competent cells. In the laboratory, bacterial cells can be made competent and DNA subsequently introduced by a procedure called the heat shock method.
Heat shock transformation uses a calcium rich environment provided by calcium chloride to counteract the electrostatic repulsion between the plasmid DNA and bacterial cellular membrane. A sudden increase in temperature creates pores in the plasma membrane of the bacteria and allows for plasmid DNA to enter the bacterial cell. This video goes through a step-by-step procedure on how to create chemically competent bacteria, perform heat shock transformation, plate the transformed bacteria, and calculate transformation efficiency.

Bacterial Transformation Using Heat Shock and Competent Cells

Transformation is the process that occurs when a cell ingests foreign DNA from its surroundings. Transformation can occur in nature in certain types of ...

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Bacterial Transformation: Electroporation

The term &#x201C;transformation&#x201D; refers cellular ingestion of foreign DNA. In nature, transformation can occur in certain types of bacteria. In molecular biology, however, transformation is artificially induced through the creation of pores in the bacterial cell walls. Bacterial cells that are able to take up DNA from the environment are called competent cells. Electrocompetent cells can be produced in the laboratory and transformation of these cells can be achieve via the application of an electrical field that creates pores in the cell wall through which DNA can pass.
The video explains the equipment used in electroporation such as an electroporator and electroporation cuvette. The video also goes through a step-by-step procedure about how to create electrocompetent cells and electroporate cells of interest. 
Prediction of the success of a transformation of an experiment, by observing the time constant, as well as the importance of removing salt from the solutions when electroporating, are also mentioned.

The term &#x201C;transformation&#x201D; refers cellular ingestion of foreign DNA.  In nature, transformation can occur in certain types of bacteria.  In ...

Transformation of E. coli Cells Using an Adapted Calcium Chloride Procedure

Source: Natalia Martin1, Andrew J. Van Alst1, Rhiannon M. LeVeque1, and Victor J. DiRita1 
1 Department of Microbiology and Molecular Genetics, Michigan State University
Bacteria have the ability to exchange genetic material (DeoxyriboNucleic Acid, DNA) in a process known as horizontal gene transfer. Incorporating exogenous DNA provides a mechanism by which bacteria can acquire new genetic traits that allow them to adapt to changing environmental conditions, such as the presence of antibiotics or antibodies (1) or molecules found in natural habitats (2). There are three mechanisms of horizontal gene transfer: transformation, transduction, and conjugation (3). Here we will focus on transformation, the ability of bacteria to take up free DNA from the environment. In the laboratory, the transformation process has four general steps: 1) Preparation of competent cells, 2) Incubation of competent cells with DNA, 3) Recovery of cells, and 4) Plating of the cells for growth of the transformants (Figure 1).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10515/10515fig1.jpg" /> 
Figure 1: General steps of the transformation process. The transformation process has four general steps: 1) Preparation of competent cells, 2) Incubation with DNA, 3) Recovery of the cells and 4) Plating cells for growth of the transformants.
For transformation to occur, the recipient bacteria must be in a state known as competence. Some bacteria have the ability to become naturally competent in response to certain environmental conditions. However, many other bacteria do not become competent naturally, or the conditions for this process are yet unknown. The ability to introduce DNA into bacteria has a range of research applications: to generate multiple copies of a DNA molecule of interest, to express large amount of proteins, as a component in cloning procedures, and others. Because of the value of transformation to molecular biology, there are several protocols aimed to make cells artificially competent when conditions for natural competence are unknown. Two main methods are used to prepare artificially competent cells: 1) through chemical treatment of cells and 2) exposing the cells to electric pulses (electroporation). The former uses different chemicals depending on the procedure to create attraction between the DNA and the cell surface, while the latter uses electric fields to generate pores in the bacterial cell membrane through which DNA molecules can enter. The most efficient approach for chemical competence is incubation with divalent cations, most notably calcium (Ca2+) (4,5) Calcium-induced competence is the procedure that will be described here (6). This method is mainly used for transformation of Gram-negative bacteria, and that will be the focus of this protocol.
The procedure of chemical transformation involves a series of steps in which cells are exposed to cations to induce chemical competence. These steps are subsequently followed by a temperature change - heat shock - that favors the uptake of foreign DNA by the competent cell (7). The bacterial cell envelopes are negatively charged. In Gram-negative bacteria like Escherichia coli, the outer membrane is negatively-charged due to the presence of lipopolysaccharide (LPS) (8). This results in repulsion of the similarly negatively-charged DNA molecules. In chemical competence induction, positively-charged calcium ions neutralize this charge repulsion enabling DNA absorbance onto the cell surface (9). Calcium treatment and incubation with DNA are done on ice. Subsequently, an incubation at higher temperatures (42&#176;C), the heat shock, is performed. This temperature imbalance further favors the DNA uptake. Bacterial cells need to be at the mid-exponential growth phase to withstand the heat shock treatment; in other growth stages the bacterial cells are too sensitive to the heat resulting in loss of viability which significantly decreases transformation efficiency.
Different DNA sources can be used for transformation. Typically, plasmids, small circular, double-stranded DNA molecules, are used for transformation in most laboratory procedures in E. coli. For plasmids to be maintained in the bacterial cell after transformation, they need to contain an origin of replication. This allows them to be replicated in the bacterial cell independently from the bacterial chromosome. Not all the bacterial cells get transformed during the transformation procedure. Thus, transformation yields a mixture of transformed cells and non-transformed cells. To distinguish between these two populations, a selection method to identify the cells that have acquired the plasmid is used. Plasmids usually contain selectable markers, which are genes encoding a trait that confers an advantage for growth (i.e. resistance to an antibiotic or chemical or rescue from a growth auxotrophy). After transformation, bacterial cells are plated on selective media, which only allows for growth of the transformed cells. In the case of cells transformed with a plasmid conferring resistance to a given antibiotic, the selective media will be growth media containing that antibiotic. Several different methods can be used to confirm that the colonies grown in the selective media are transformants (i.e. have incorporated the plasmid). For example, plasmids can be recovered from these cells using plasmid preparation methods (10) and digested to confirm plasmid size. Alternatively, colony PCR can be used to confirm the presence of the plasmid of interest (11).
The aim of this experiment is to prepare E. coli DH5&#945; chemically competent cells, using an adaptation of the calcium chloride procedure (12), and to transform them with the plasmid pUC19 to determine transformation efficiency. The E. coli strain DH5&#945; is a commonly used strain in molecular biology applications. Because of its genotype, specifically the recA1 and endA1, this strain allows for increased insert stability and improve the quality of plasmid DNA in subsequent preparations. Since the transformation efficiency decreases with increasing size of the DNA, the plasmid pUC19 was used in this protocol because of its small size (2686 bp) (see https://www.mobitec.com/cms/products/bio/04_vector_sys/standard_cloning_vectors.html for a vector map). pUC19 confers resistance to ampicillin and thus, this was the antibiotic used for selection.

Transformation of E. coli: Adapted Calcium Chloride Procedure

Source: Natalia Martin1, Andrew J. Van Alst1, Rhiannon M. LeVeque1, and Victor J. DiRita1
1 Department of Microbiology and Molecular Genetics, Michigan ...

Methods for Electroporation and Transformation Confirmation in Limosilactobacillus reuteri DSM20016

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Rapid Protocol for Preparation of Electrocompetent Escherichia coli and Vibrio cholerae

Generation of Null Mutants to Elucidate the Role of Bacterial Glycosyltransferases in Bacterial Motility

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Bacterial Transformation: The Heat Shock Method

Bacterial Transformation: Electroporation

Transformation of E. coli Cells Using an Adapted Calcium Chloride Procedure