We would like to thank the Canadian Foundation for Innovation, the British Columbia Knowledge Development Fund, and the Canadian Institutes of Health Research (CIHR) for supporting this work. M.T was supported by a fellowship from the TULA foundation funded Centre for Microbial Diversity and Evolution (CMDE).

Expression of recombinant proteins in the methylotrophic yeast Pichia pastoris
Before you start this protocol you should have your gene of interest cloned in frame in a P. pastoris parent vector and have it sequenced to check for the correct insertion of the gene of interest in the vector. 
Step I: Generating electrocompetent yeast cells, linearization of the construct and transformation into P. pastoris
For this step you will need to have the following media and plates at hand:
<ul>
 <li>YPDS plates </li>
 <li>600 mL YPD medium </li>
 <li>ice-cold sterile water </li>
 <li>1 M sorbitol </li>
 <li>Amicon Ultra-4 Centrifugal Filter Device</li>
 <li>Microcon YM-30 Centrifugal Filter Unit </li>
 <li>0.2 cm electroporation cuvette </li>
 <li>15 mL sterile glass tube </li>
 <li>sterile glass Pasteur Pipette </li>
 <li>YPDS plates containing Zeocin </li>
</ul>
Part 1: Preparation of electrocompetent yeast cells
<ol>
	<li>Four days before the intended transformation streak out P. pastoris cells on a YPDS plate without Zeocin and let them grow a 30&deg;C for 1-2 days or until single colonies form.</li>
	<li>Two days before intended transformation grow 5 ml of your P. pastoris strain in YPD medium in a 50 ml Falcon tube at 30&deg;C overnight. Place the Falcon tube in a flask to fix it in the shaker.</li>
	<li>The day before the transformation inoculate 500 mL of fresh YPD medium in a 2 liter flask with 0.25 mL of the overnight culture and let grow overnight again in a shaker at 30&deg;C to an OD600 = 1.3-1.5. 
 Note: dilute your sample before measuring on the spectrophotometer so that you get an accurate result.</li>
	<li>The day of the transformation have ice-cold sterile water and 1 M sorbitol at hand. Centrifuge the cells at 1500 x g for 5 minutes at 4&deg;C. Resuspend the pellet with 500 mL of ice-cold, sterile water.</li>
	<li>Centrifuge the cells again as step 4. Resuspend the pellet with 250 mL of ice-cold, sterile water.</li>
	<li>Centrifuge the cells again as in step 4. Resuspend the pellet in 20 mL of ice-cold 1 M sorbitol.</li>
	<li>Centrifuge the cells again as in step 4. Resuspend the pellet in 1 mL of ice-cold 1 M sorbitol to a final volume of ~ 1.5 mL. Store cells on ice or at 4&deg;C until further use. 
 Note: we prepare more electrocompetent cells than needed and store them in aliquots of 80 &mu;l in microcentrifuge tubes at -80 &deg;C. We use the stored cells for no longer than one month.</li>
</ol>
Part 2: Linearization and concentration of the pPICZ&alpha;A construct
For transformation linearize the vector containing your gene of interest by restriction digest. We use the enzyme PmeI. Other restriction sites are possible. You have to make sure that your insert does not contain the restriction site you want to use to linearize your vector. For transformation into P. pastoris you will need 5-20 &mu;g of linearized DNA in 5-10 &mu;l sterile water.
Also linearize, concentrate and transfer the plain vector (no insert) into P. pastoris. The parent vector without the insert is a control for background intracellular expression and allows you to interpret your expression results. 
<ol>
	<li>Thaw all reagents on ice, combine the following reagents and briefly centrifuge the tube to get all liquid to the bottom, tap the tube and spin again:
<ul>
 <li>x &mu;l sterile water</li>
 <li>5.0 &mu;l 10X NEBuffer 4</li>
 <li>0.5 &mu;l 100X BSA</li>
 <li>up to 2 &mu;g vector DNA (~ 100 ng/&mu;l)</li>
 <li>2.0 &mu;l PmeI enzyme mix</li>
 </ul>
 &rarr;50 &mu;l total reaction volume Note: to obtain sufficient linearized vector DNA prepare the above mix 3 or 4 times in separate tubes and combine the solutions when concentrating the digested DNA via Amicon Ultra Centrifugal Device.</li>
	<li>Incubate at 37&deg;C for 3 hours and heat inactivate the enzyme mix at 65&deg;C for 20 minutes.</li>
	<li>In the mean time pre-rinse an Amicon Ultra-4 Centrifugal Filter Device with 1 mL of Milli-Q &nbsp;water, spin the tube at 4000 x g for 6-8 minutes and discard the flow through. Note: do not allow the membrane to dry out once wet. If you are not using the device after pre-rinsing, leave water on the membrane until the device is used.</li>
	<li>Transfer the heat-inactivated solution from step 2 to the pre-rinsed Amicon Centrifugal Filter Unit, spin the tube at 4000 x g for 6-8 minutes and discard the flow through. Note: if you have prepared more than one linearization mix, combine all solutions into one Amicon Centrifugal Filter Unit.</li>
	<li>Centrifuge until the amount of solution on the Amicon filter is reduced to approximately 100 ~ 150 &mu;l. Add another 1.5 mL of sterile water to the empty tube from above and transfer the solution to the Amicon tube and repeat the centrifugation step. Wash a second time with 1.5 mL of sterile water and discard flow through again. Reduce the final volume to ~ 150 &mu;l. The washing steps remove the remaining salt from the buffer to prevent arching when pulsing the cells. Note: we use a swinging bucket rotor for the centrifugation step; simply place the capped Amicon centrifugal filter unit in a 50 mL Falcon tube to hold it in place during centrifugation. Amicon filter devices retain 50 &mu;l of solution on the filter even after extended centrifugation.</li>
	<li>For further concentration of the DNA preparation transfer the remaining DNA solution from the Amicon filter device to a pre-rinsed Microcon YM-30 Centrifugal Filter Unit and rinse the Amicon tube with an additional 50 &mu;l of sterile water to recover all DNA. Centrifuge the Microcon Centrifugal Filter Unit in a microcentrifuge at 10,000 x g until the filter is still slightly covered with liquid. Only spin for a short time and check every minute if only a small amount of liquid is left on the filter. To recover your DNA solution, place the filter upside down in a new microcentrifuge tube and spin in a second centrifugation step at 1000 x g for 3 minutes. The final volume should not exceed 10-15 &mu;l in total, otherwise your DNA might be too diluted for the transformation step. Note: do not spin the filter device too long and let it not dry completely to prevent potential sample loss. If your membrane is dry, add 10-15 &mu;l sterile water onto the membrane, agitate gently for 30 seconds and recover your DNA as described above.</li>
</ol>
Part 3: Transformation into P. pastoris by electroporation
<ol>
	<li>The linearized and concentrated pPICZ&alpha;A DNA containing your insert is now ready for transformation into the electrocompetent P. pastoris cells (see step I, part 2). About 15 minutes before the transformation prepare the following reagents and devices: fill a microcentrifuge tube with 1 mL of 1 M sorbitol and place it on ice; place a 0.2 cm electroporation cuvette on ice; label a sterile 15 mL glass tube and have a sterile glass Pasteur pipette at hand.</li>
	<li>Transfer 80 &mu;l of the cells from step I, part 2.7 to an ice-cold 0.2 cm electroporation cuvette. Add the concentrated linearized pPICZ&alpha;A DNA solution from step I, part 3.6 and mix by moving the pipette tip from side-to-side in the electroporation cuvette. Note: when adding the DNA only push pipette to the first stop. After mixing the DNA with the cells push pipette to the second stop and slowly remove from the cuvette.</li>
	<li>Incubate the cuvette with the cells on ice for 5 minutes.</li>
	<li>Wipe the outside of the cuvette with a tissue and pulse the cells according to the parameters for yeast (Saccharomyces cerevisiae) as suggested by the manufacturer of the specific electroporation device being used. Note: we use a Bio-Rad GenePulser with the following conditions:
<ul>
 <li>Charging voltage (V): 1500;</li>
 <li>Capacitance (&mu;F): 25;</li>
 <li>Resistance (&Omega;): 200. </li>
</ul>
Using a 0.2 cm electroporation cuvette, the protocol generates a pulse length of ~10 ms with a field strength of ~7500 V/cm.</li>
	<li>Immediately add 1 mL of ice-cold 1 M sorbitol to the cuvette. Transfer the cuvette content to a sterile 15 mL tube using the sterile glass Pasteur pipette.</li>
	<li>Let the tube incubate at 30&deg;C without shaking for 1.5 hours.</li>
	<li>Add 5-7 sterilized glass beads on four labeled YPDS plates containing 100 &mu;g/mL Zeocin. Spread 250 &#956;l of the electroporation mix from the 15 mL glass tube on each plate. Shake the plate horizontally to evenly spread out the cells. Let the plate dry for 15 minutes and then remove the beads from the agar by inverting the plate. Note: we use 100 &mu;g/mL Zeocin to select for transformants when using the GS115 Pichia strain. The selection conditions may vary if you are using another Pichia strain.</li>
	<li>Incubate plates upside down for 2 to 3 days at 30&deg;C until colonies form. Wrap the plates in a black plastic to prevent degradation of the light sensitive Zeocin. Less amount of antibiotic in the plates can result in false positive clones.</li>
	<li>After colonies have formed, pick 12 colonies and purify them by streaking the clones on fresh YPDS plates containing 100 &mu;g/mL of Zeocin.</li>
</ol>
Step II: Protein expression in Pichia pastoris
For this step you will need to have the following media, plates and reagents at hand:
<ul>
 <li>BMGY medium</li>
 <li>glycerol, sterilized</li>
 <li>BMMY medium</li>
 <li>methanol, sterilized</li>
 <li>EPICENTRE Yeast DNA purification kit</li>
</ul>
You also need the following flasks:
<ul>
 <li>250 mL flask, autoclaved</li>
 <li>1 L baffled flask, autoclaved</li>
 <li>200 mL beaker, autoclaved</li>
</ul>
Part 1: Protein expression in Pichia pastoris
All of the following steps are also performed for the transformed control vector (with no insert).
<ol>
	<li>Pick a single colony from the purified Pichia colonies from step I, part 3.9 and inoculate in 25 mL BMGY in a sterile 250 mL flask. Grow at 30&deg;C in a shaking incubator (250-300 rpm) until culture reaches an OD600 = 2-6 (approximately 16-18 hours). Note: dilute your sample before measuring on the spectrophotometer so that you get an accurate result. The cells will be in log-phase growth.</li>
	<li>When the cells have reached the appropriate OD600 prepare a glycerol stock. Transfer 800 &mu;l of the 25 mL cell culture to a 2 mL Corning cyrogenic vial and add 200 &mu;l of sterilized glycerol. Freeze the glycerol stock and store at -80&deg;C.</li>
	<li>For yeast DNA purification, transfer 1.5 mL of the 25 mL cell culture to a microcentrifuge tube. Harvest the cells by centrifuging at 1300 x g for 1 minute at room temperature. These cells are used for analyzing Pichia integrants to double check if the gene of interest has integrated into the Pichia genome (see step II, part2). Store the cell pellet at 4&deg;C until further analysis.</li>
	<li>Transfer the rest of the 25 mL culture to a 50 mL Falcon tube and harvest the cells by centrifuging at 3000 x g for 5 minutes at room temperature. Decant supernatant and place the tubes upside down on a tissue to remove any residual media. To wash the cell pellet resuspend it in 20 mL BMMY to remove any remaining BMGY medium, as glycerol from the BMGY medium can inhibit the later expression. Centrifuge again at 3000 x g for 5 minutes at room temperature and decant the supernatant.</li>
	<li>Resuspend cell pellet to an OD600 of 1.0 in BMMY medium to induce expression (approximately 100-200 mL) and transfer the culture in a 1 liter baffled flask. Cover the flask with a 200 mL beaker and return to incubator to continue growth at 30&deg;C. Note: it is important that the temperature does not exceed 30&deg;C. If the temperature of your incubator fluctuates, set the temperature at 28&deg;C. Adequate aeration is also an important parameter for efficient expression during methanol induction (when inducing expression, never exceed a culture volume &gt; 10-30% of your total flask volume). We highly recommend the use of baffled flasks, as they introduce more oxygen in the culture media.</li>
	<li>Add sterilized pure methanol to a final concentration of 0.5% methanol every 24 hours to maintain induction.</li>
	<li>At certain time points after the start of the expression transfer 1 mL of the culture to a 1.5 mL microcentrifuge tube. Centrifuge at 1300 x g for 2.5 minutes at room temperature. Transfer the supernatant to a separate 1.5 mL microcentrifuge tube. Store the supernatant and cell pellets at -80&deg;C until further analysis. The samples from different time points will be analyzed to establish the optimal time period for protein expression after induction. Note: we choose 6h, 12h, 24h, 36h and 48h as the time points for taking the samples. Further growth up to 4 days is possible. The optimal time points vary between different expressed proteins.</li>
	<li>Analyze the supernatants and cell pellets for protein expression by Coomassie-stained SDS-PAGE and western blot (not described in this protocol).</li>
</ol>
Part 2: Yeast DNA purification and PCR analysis of the Pichia integrant
All reagents necessary for the yeast DNA purification step of the Pichia culture in BMGY are included in the MasterPure Yeast DNA Purification Kit from EPICENTRE. Note: this analysis is additional and is performed to determine if the gene of interest has integrated into the Pichia genome. The cell pellet from a 1.5 mL sample taken from the culture in BMGY (see step II, part 1.3) is used.
Follow the instructions of the Epicentre MasterPure Yeast DNA Purification manual and run a PCR with the yeast DNA with specific primers for the insert. Analyze 3 &mu;l of your PCR product on an agarose gel containing 1 &mu;l ethidium bromide. Include a 1 Kb+ size marker in one well to interpret the size of your insert and visualize the DNA using a gel documentation station.
The recombinant protein can be analyzed by western blot analysis. Protein purification can be performed by His-tag purification on metal-charged resin (not described in this protocol).
Appendix, list of recipes
Step I:
<ol>
	<li>YPD medium: To prepare 600 mL of YPD medium (Yeast Extract Peptone Dextrose Medium) dissolve 5 g yeast extract and 12 g peptone in 540 mL of water. Autoclave for 20 minutes on liquid cycle. In the mean time prepare 70 mL of 20% dextrose and filter-sterilize before use. Let the autoclaved solution cool to ~60&deg;C and add 60 mL of filter-sterilized 20% dextrose.</li>
	<li>YPDS (+ Zeocin) plates: To prepare 500 mL of YPDS + Zeocin agar (Yeast Extract Peptone Dextrose Medium with Sorbitol) dissolve 5 g yeast extract, 91.1 g sorbitol and 10 g peptone in 450 mL of water. Add 10 g of agar and a magnetic stir bar and autoclave for 20 minutes on liquid cycle. In the mean time prepare 60 mL of 20% dextrose and filter-sterilize before use. Let the autoclaved solution cool to ~ 60&deg;C, add 50 mL of filter-sterilized 20% dextrose. Pour a few plates before adding the antibiotic for YPDS plates without Zeocin. Then add 500 &mu;l Zeocin from a 100 mg/ml stock solution to obtain a final concentration of 100 &mu;g/ml Zeocin in the agar. Let stir on a magnetic plate when adding the antibiotic and let stir for about an additional 2 minutes to equally mix. Pour media into Petri dishes and cover with black plastic. Let plates dry on bench overnight. Store YPD plates containing Zeocin at 4&deg;C. The shelf life is one to two weeks. Note: the coverage with the plastic is necessary because Zeocin is light sensitive.</li>
</ol>
Step II:
<ol>
	<li>BMGY (Buffered Glycerol-complex Medium) and BMMY (Buffered Methanol-complex Medium): Dissolve for each medium 8 g of yeast extract and 16 g peptone in 560 mL water. Autoclave for 20 minutes on liquid cycle. In the mean time prepare the following solutions:
<ul>
 <li>1 M potassium phosphate buffer, pH 6.0:</li>
</ul>
Combine 24 mL of 1M K2HPO4 and 156 mL of 1M K2HPO4 and confirm that the pH = 6.0 (if the pH needs to be adjusted, use phosphoric acid or KOH). Filter sterilize and store at room temperature. The shelf life of this solution is greater than one year.
<ul>
 <li>10X YNB (13.4% Yeast Nitrogen Base with Ammonium Sulfate without amino acids): </li>
</ul>
Dissolve 26.8 g of yeast nitrogen base (YNB) with ammonium sulfate and without amino acids in 200 mL of water and filter sterilize. Heat the solution to dissolve YNB completely in water. Store at 4&deg;C. The shelf life of this solution is approximately one year. Note: alternatively, use 3.4 g of YNB without ammonium sulfate and without amino acids and add 10 g of ammonium sulfate.
<ul>
 <li>500X B (0.02% Biotin): </li>
</ul>
Dissolve 10 mg biotin in 50 mL of water and filter sterilize. Store at 4&deg;C. The shelf life of this solution is approximately one year.
<ul>
 <li>10X M (5% Methanol): </li>
</ul>
Mix 5 mL of methanol with 95 mL of water. Filter sterilize and store at 4&deg;C. The shelf life of this solution is approximately two months.
<ul>
 <li>10X GY (10% Glycerol):</li>
</ul>
Mix 10 mL of glycerol with 90 mL of water. Filter sterilize. Store at room temperature. The shelf life of this solution is greater than one year. Let the autoclaved solution cool to room temperature, then add the following and mix well:
<ul>
 <li>80 mL 1 M potassium phosphate buffer, pH 6.0; </li>
 <li>80 mL 10X YNB</li>
 <li>0.16 mL 500X B</li>
 <li>80 mL 10X GY</li>
</ul>
For preparation of BMMY, add 80 mL 10X M instead of glycerol. 
Store media at 4&deg;C. The shelf life of this solution is approximately two months.</li></ol>
Representative Results
<img src="/files/ftp_upload/1862/1862fig1.jpg" alt="Figure 1" /> Figure 1. This western blot image shows four expressed proteins (labeled as 1605, 0537, 1228, and 1682) after 24 hours of expression. The antibody used for immunoblotting was a c-myc-HRP antibody. The second lane (labeled N) is a negative control and was loaded with supernatant from cells that do not express a recombinant protein because they were transformed with the parent (plain) vector.

<ol>
	<li>Cereghino, G. P., Cregg, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Applications+of+yeast+in+biotechnology:+protein+production+and+genetic+analysis.">Applications of yeast in biotechnology: protein production and genetic analysis.</a> Curr Opin Biotechnol. 10, 422-427 (1999).</li><li>Cregg, J. M., Barringer, K. J., Hessler, A. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pichia+pastoris+as+a+Host+System+for+Transformations.">Pichia pastoris as a Host System for Transformations.</a> Mol. Cell. Biol. 5, 3376-3385 (1985).</li><li>Cregg, J. M., Cereghino, J. L., Shi, J., Higgins, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recombinant+protein+expression+in+Pichia+pastoris.">Recombinant protein expression in Pichia pastoris.</a> Mol Biotechnol. 16, 23-52 (2000).</li><li>Cereghino, G. P., Cereghino, J. L., Ilgen, C., Cregg, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Production+of+recombinant+proteins+in+fermenter+cultures+of+the+yeast+Pichia+pastoris.">Production of recombinant proteins in fermenter cultures of the yeast Pichia pastoris.</a> Curr Opin Biotechnol. 4, 329-332 (2002).</li></ol>

Protein expression using the methylotrophic yeast Pichia pastoris as a host system is described in this protocol. The protein can be secreted into the medium depending on the used cloning vector. The secretion of the recombinant protein makes the subsequent purification easier. However, the shown conditions may have to be optimized for the expression of different proteins and changes in the conditions and expression times may result in increased protein levels. The vector used in this protocol has the feature of...

expression of recombinant proteins in the methylotrophic yeast pichia pastoris

Protein expression in the microbial eukaryotic host Pichia pastoris offers the possibility to generate high amounts of recombinant protein in a fast and easy to use expression system.
As a single-celled microorganism P. pastoris is easy to manipulate and grows rapidly on inexpensive media at high cell densities. Being a eukaryote, P. pastoris is able to perform many of the post-translational modifications performed by higher eukaryotic cells and the obtained recombinant proteins undergo protein folding, proteolytic processing, disulfide bond formation and glycosylation [1].
As a methylotrophic yeast P. pastoris is capable of metabolizing methanol as its sole carbon source. The strong promoter for alcohol oxidase, AOX1, is tightly regulated and induced by methanol and it is used for the expression of the gene of interest. Accordingly, the expression of the foreign protein can be induced by adding methanol to the growth medium [2; 3].
Another important advantage is the secretion of the recombinant protein into the growth medium, using a signal sequence to target the foreign protein to the secretory pathway of P. pastoris. With only low levels of endogenous protein secreted to the media by the yeast itself and no added proteins to the media, a heterologous protein builds the majority of the total protein in the medium and facilitates following protein purification steps [3; 4].
The vector used here (pPICZ&alpha;A) contains the AOX1 promoter for tightly regulated, methanol-induced expression of the gene of interest; the &alpha;-factor secretion signal for secretion of the recombinant protein, a Zeocin resistance gene for selection in both E. coli and Pichia and a C-terminal peptide containing the c-myc epitope and a polyhistidine (6xHis) tag for detection and purification of a recombinant protein. We also show western blot analysis of the recombinant protein using the specific Anti-myc-HRP antibody recognizing the c-myc epitope on the parent vector.

Watch this Scientific Journal Video about Expression of Recombinant Proteins in the Methylotrophic Yeast Pichia pastoris at JoVE.com

Expression of Recombinant Proteins in the Methylotrophic Yeast Pichia pastoris

The protocol describes protein expression using the methylotrophic yeast Pichia pastoris. The preparation of electrocompetent yeast cells, transformation of the vector with the gene of interest into P. pastoris and yeast DNA purification are also performed. Western blot analysis and protein purification build the last steps in this protein expression protocol.

Expression of Recombinant Proteins in the Methylotrophic Yeast Pichia pastoris

Protein expression in the microbial eukaryotic host Pichia pastoris offers the possibility to generate high amounts of recombinant protein in a fast and ...

expression-recombinant-proteins-methylotrophic-yeast-pichia

Research

JoVE Journal

Biology

45.6K Views.  University of British Columbia - UBC. The protocol describes protein expression using the methylotrophic yeast Pichia pastoris. The preparation of electrocompetent yeast cells, transformation of the vector with the gene of interest into P. pastoris and yeast DNA purification are also performed. Western blot analysis and protein purification build the last steps in this protein expression protocol. 

Video: Expression of Recombinant Proteins in the Methylotrophic Yeast Pichia pastoris

Staining of Proteins in Gels with Coomassie G-250 without Organic Solvent and Acetic Acid

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents (Methanol, Ethanol or 2-Propanol) and acetic acid are used for fixation, staining and destaining of proteins in a gel after SDS-PAGE. To speed up the procedure, heating the staining solution in the microwave oven for a short time is frequently used. This usually results in evaporation of toxic or hazardous Methanol, Ethanol or 2-Propanol and a strong smell of acetic acid in the lab which should be avoided due to safety considerations. In a protocol originally published in two patent applications by E.M. Wondrak (US2001046709 (A1), US6319720 (B1)), an alternative composition of the staining solution is described in which no organic solvent or acid is used. The CBB is dissolved in bidistilled water (60-80mg of CBB G-250 per liter) and 35 mM HCl is added as the only other compound in the staining solution. The CBB staning of the gel is done after SDS-PAGE and thorough washing of the gel in bidistilled water. By heating the gel during the washing and staining steps, the process can be finished faster and no toxic or hazardous compunds are evaporating. The staining of proteins occurs already within 1 minute after heating the gel in staining solution and is fully developed after 15-30 min with a slightly blue background that is destained completely by prolonged washing of the stained gel in bidistilled water, without affecting the stained protein bands.

A short protocol for protein staining with Coomassie Brilliant Blue (CBB) G-250 in polyacrylamide gels is described without using organic solvents or acetic acid as in the classical staining procedures with CBB.

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents ...

Using an Automated Cell Counter to Simplify Gene Expression Studies: siRNA Knockdown of IL-4 Dependent Gene Expression in Namalwa Cells

The use of siRNA mediated gene knockdown is continuing to be an important tool in studies of gene expression.  siRNA studies are being conducted not only to study the effects of downregulating single genes, but also to interrogate signaling pathways and other complex interaction networks.  These pathway analyses require both the use of relevant cellular models and methods that cause less perturbation to the cellular physiology.  Electroporation is increasingly being used as an effective way to introduce siRNA and other nucleic acids into difficult to transfect cell lines and primary cells without altering the signaling pathway under investigation. There are multiple critical steps to a successful siRNA experiment, and there are ways to simplify the work while improving the data quality at several experimental stages.  To help you get started with your siRNA mediated gene knockdown project, we will demonstrate how to perform a pathway study complete from collecting and counting the cells prior to electroporation through post transfection real-time PCR gene expression analysis.  The following study investigates the role of the transcriptional activator STAT6 in IL-4 dependent gene expression of CCL17 in a Burkitt lymphoma cell line (Namalwa).  The techniques demonstrated are useful for a wide range of siRNA-based experiments on both adherent and suspension cells.  We will also show how to streamline cell counting with the TC10 automated cell counter, how to electroporate multiple samples simultaneously using the MXcell electroporation system, and how to simultaneously assess RNA quality and quantity with the Experion automated electrophoresis system.

This procedure describes a quick and easy workflow to introduce siRNA into difficult to transfect cell lines and follow gene expression by real-time PCR. Use of an automated cell counter, multi-well electroporation plate, and automated electrophoresis station provide quick and reliable results without the need for expensive robotic handling.

The use of siRNA mediated gene knockdown is continuing to be an important tool in studies of gene expression.  siRNA studies are being conducted not only ...

In vitro Reconstitution of the Active T. castaneum Telomerase

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more than a decade. Here, we present methods for the isolation of the recombinant Tribolium castaneum TERT (TcTERT) and the reconstitution of the active T. castaneum telomerase ribonucleoprotein (RNP) complex in vitro.
Telomerase is a specialized reverse transcriptase1 that adds short DNA repeats, called telomeres, to the 3' end of linear chromosomes2 that serve to protect them from end-to-end fusion and degradation. Following DNA replication, a short segment is lost at the end of the chromosome3 and without telomerase, cells continue dividing until eventually reaching their Hayflick Limit4. Additionally, telomerase is dormant in most somatic cells5 in adults, but is active in cancer cells6 where it promotes cell immortality7.
The minimal telomerase enzyme consists of two core components: the protein subunit (TERT), which comprises the catalytic subunit of the enzyme and an integral RNA component (TER), which contains the template TERT uses to synthesize telomeres8,9. Prior to 2008, only structures for individual telomerase domains had been solved10,11. A major breakthrough in this field came from the determination of the crystal structure of the active12, catalytic subunit of T. castaneum telomerase, TcTERT1.
Here, we present methods for producing large quantities of the active, soluble TcTERT for structural and biochemical studies, and the reconstitution of the telomerase RNP complex in vitro for telomerase activity assays. An overview of the experimental methods used is shown in Figure 1.

In vitro Reconstitution of the Active T. castaneum Telomerase

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more than a decade. Here, we present methods for the isolation of the recombinant Tribolium castaneum TERT (TcTERT) and the reconstitution of the active T. castaneum telomerase ribonucleoprotein (RNP) complex in vitro.

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more ...

High-throughput Purification of Affinity-tagged Recombinant Proteins

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further biochemical analyses to obtain detailed insights into structure/function relationships. Advances in oligonucleotide- and gene synthesis technology make large-scale mutagenesis strategies increasingly feasible, including the substitution of target residues by all 19 other amino acids. Gain- or loss-of-function phenotypes then allow systematic conclusions to be drawn, such as the contribution of particular residues to catalytic activity, protein stability and/or protein-protein interaction specificity.
In order to attribute the different phenotypes to the nature of the mutation - rather than to fluctuating experimental conditions - it is vital to purify and analyse the proteins in a controlled and reproducible manner. High-throughput strategies and the automation of manual protocols on robotic liquid-handling platforms have created opportunities to perform such complex molecular biological procedures with little human intervention and minimal error rates1-5.
Here, we present a general method for the purification of His-tagged recombinant proteins in a high-throughput manner. In a recent study, we applied this method to a detailed structure-function investigation of TFIIB, a component of the basal transcription machinery. TFIIB is indispensable for promoter-directed transcription in vitro and is essential for the recruitment of RNA polymerase into a preinitiation complex6-8. TFIIB contains a flexible linker domain that penetrates the active site cleft of RNA polymerase9-11. This linker domain confers two biochemically quantifiable activities on TFIIB, namely (i) the stimulation of the catalytic activity during the 'abortive' stage of transcript initiation, and (ii) an additional contribution to the specific recruitment of RNA polymerase into the preinitiation complex4,5,12 . We exploited the high-throughput purification method to generate single, double and triple substitution and deletions mutations within the TFIIB linker and to subsequently analyse them in functional assays for their stimulation effect on the catalytic activity of RNA polymerase4. Altogether, we generated, purified and analysed 381 mutants - a task which would have been time-consuming and laborious to perform manually. We produced and assayed the proteins in multiplicates which allowed us to appreciate any experimental variations and gave us a clear idea of the reproducibility of our results.
This method serves as a generic protocol for the purification of His-tagged proteins and has been successfully used to purify other recombinant proteins. It is currently optimised for the purification of 24 proteins but can be adapted to purify up to 96 proteins.

We describe a method for the affinity-tagged purification of recombinant proteins using liquid-handling robotics. This method is generally applicable to the small-scale purification of soluble His-tagged proteins in a high-throughput format.

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further ...

Efficient Agroinfiltration of Plants for High-level Transient Expression of Recombinant Proteins

Mammalian cell culture is the major platform for commercial production of human vaccines and therapeutic proteins. However, it cannot meet the increasing worldwide demand for pharmaceuticals due to its limited scalability and high cost. Plants have shown to be one of the most promising alternative pharmaceutical production platforms that are robust, scalable, low-cost and safe. The recent development of virus-based vectors has allowed rapid and high-level transient expression of recombinant proteins in plants. To further optimize the utility of the transient expression system, we demonstrate a simple, efficient and scalable methodology to introduce target-gene containing Agrobacterium into plant tissue in this study. Our results indicate that agroinfiltration with both syringe and vacuum methods have resulted in the efficient introduction of Agrobacterium into leaves and robust production of two fluorescent proteins; GFP and DsRed. Furthermore, we demonstrate the unique advantages offered by both methods. Syringe infiltration is simple and does not need expensive equipment. It also allows the flexibility to either infiltrate the entire leave with one target gene, or to introduce genes of multiple targets on one leaf. Thus, it can be used for laboratory scale expression of recombinant proteins as well as for comparing different proteins or vectors for yield or expression kinetics. The simplicity of syringe infiltration also suggests its utility in high school and college education for the subject of biotechnology. In contrast, vacuum infiltration is more robust and can be scaled-up for commercial manufacture of pharmaceutical proteins. It also offers the advantage of being able to agroinfiltrate plant species that are not amenable for syringe infiltration such as lettuce and Arabidopsis. Overall, the combination of syringe and vacuum agroinfiltration provides researchers and educators a simple, efficient, and robust methodology for transient protein expression. It will greatly facilitate the development of pharmaceutical proteins and promote science education.

Plants offer a novel system for the production of pharmaceutical proteins on a commercial scale that is more scalable, cost-efficient and safe than current expression paradigms. In this study, we report a simple and convenient, yet scalable approach to introduce target-gene containing Agrobacterium tumefaciens into plants for protein transient expression.

Mammalian cell culture is the major platform for commercial production of human vaccines and therapeutic proteins. However, it cannot meet the increasing ...

Temporal Quantification of MAPK Induced Expression in Single Yeast Cells

The quantification of gene expression at the single cell level uncovers novel regulatory mechanisms obscured in measurements performed at the population level. Two methods based on microscopy and flow cytometry are presented to demonstrate how such data can be acquired. The expression of a fluorescent reporter induced upon activation of the high osmolarity glycerol MAPK pathway in yeast is used as an example. The specific advantages of each method are highlighted. Flow cytometry measures a large number of cells (10,000) and provides a direct measure of the dynamics of protein expression independent of the slow maturation kinetics of the fluorescent protein. Imaging of living cells by microscopy is by contrast limited to the measurement of the matured form of the reporter in fewer cells. However, the data sets generated by this technique can be extremely rich thanks to the combinations of multiple reporters and to the spatial and temporal information obtained from individual cells. The combination of these two measurement methods can deliver new insights on the regulation of protein expression by signaling pathways.

Two complementary methods based on flow cytometry and microscopy are presented which enable the quantification, at the single cell level, of the dynamics of gene expression induced by the activation of a MAPK pathway in yeast.

The quantification of gene expression at the  single cell level uncovers novel regulatory mechanisms obscured in measurements  performed at the population ...

High Throughput Quantitative Expression Screening and Purification Applied to Recombinant Disulfide-rich Venom Proteins Produced in E. coli

Escherichia coli (E. coli) is the most widely used expression system for the production of recombinant proteins for structural and functional studies. However, purifying proteins is sometimes challenging since many proteins are expressed in an insoluble form. When working with difficult or multiple targets it is therefore recommended to use high throughput (HTP) protein expression screening on a small scale (1-4&#160;ml cultures) to quickly identify conditions for soluble expression. To cope with the various structural genomics programs of the lab, a quantitative (within a range of 0.1-100 mg/L culture of recombinant protein) and HTP protein expression screening protocol was implemented and validated on thousands of proteins. The protocols were automated with the use of a liquid handling robot but can also be performed manually without specialized equipment.
Disulfide-rich venom proteins are gaining increasing recognition for their potential as therapeutic drug leads. They can be highly potent and selective, but their complex disulfide bond networks make them challenging to produce. As a member of the FP7 European Venomics project (<a href="http://www.venomics.eu">www.venomics.eu</a>), our challenge is to develop successful production strategies with the aim of producing thousands of novel venom proteins for functional characterization. Aided by the redox properties of disulfide bond isomerase DsbC, we adapted our HTP production pipeline for the expression of oxidized, functional venom peptides in the E. coli cytoplasm. The protocols are also applicable to the production of diverse disulfide-rich proteins. Here we demonstrate our pipeline applied to the production of animal venom proteins. With the protocols described herein it is likely that soluble disulfide-rich proteins will be obtained in as little as a week. Even from a small scale, there is the potential to use the purified proteins for validating the oxidation state by mass spectrometry, for characterization in pilot studies, or for sensitive micro-assays.

High Throughput Quantitative Expression Screening and Purification Applied to Recombinant Disulfide-rich Venom Proteins Produced in E. coli

A protocol for the quantitative, high throughput expression screening and analytical purification of fusion proteins from small-scale Escherichia coli cultures is described and applied to the expression of disulfide-rich animal venom protein targets.

Escherichia coli (E. coli) is the most widely used expression system for the production of recombinant proteins for structural and functional studies. ...

Transient Expression of Proteins by Hydrodynamic Gene Delivery in Mice

Efficient expression of transgenes in vivo is of critical importance in studying gene function and developing treatments for diseases. Over the past years, hydrodynamic gene delivery (HGD) has emerged as a simple, fast, safe and effective method for delivering transgenes into rodents. This technique relies on the force generated by the rapid injection of a large volume of physiological solution to increase the permeability of cell membranes of perfused organs and thus deliver DNA into cells. One of the main advantages of HGD is the ability to introduce transgenes into mammalian cells using naked plasmid DNA (pDNA). Introducing an exogenous gene using a plasmid is minimally laborious, highly efficient and, contrary to viral carriers, remarkably safe. HGD was initially used to deliver genes into mice, it is now used to deliver a wide range of substances, including oligonucleotides, artificial chromosomes, RNA, proteins and small molecules into mice, rats and, to a limited degree, other animals. This protocol describes HGD in mice and focuses on three key aspects of the method that are critical to performing the procedure successfully: correct insertion of the needle into the vein, the volume of injection and the speed of delivery. Examples are given to show the application of this method to the transient expression of two genes that encode secreted, primate-specific proteins, apolipoprotein L-I (APOL-I) and haptoglobin-related protein (HPR).

In vivo transfection of naked DNA by hydrodynamic gene delivery introduces genes into the tissue of an animal with minimal inflammatory response. Sufficient amounts of gene product are generated such that gene function and regulation as well as protein structure and function can be analyzed.

Efficient expression of transgenes in vivo is of critical importance in studying gene function and developing treatments for diseases. Over the past ...

High Yield Expression of Recombinant Human Proteins with the Transient Transfection of HEK293 Cells in Suspension

The art of producing recombinant proteins with complex post-translational modifications represents a major challenge for studies of structure and function. The rapid establishment and high recovery from transiently-transfected mammalian cell lines addresses this barrier and is an effective means of expressing proteins that are naturally channeled through the ER and Golgi-mediated secretory pathway. Here is one protocol for protein expression using the human HEK293F and HEK293S cell lines transfected with a mammalian expression vector designed for high protein yields. The applicability of this system is demonstrated using three representative glycoproteins that expressed with yields between 95-120 mg of purified protein recovered per liter of culture. These proteins are the human Fc&#947;RIIIa and the rat &#945;2-6 sialyltransferase, ST6GalI, both expressed with an N-terminal GFP fusion, as well as the unmodified human immunoglobulin G1 Fc. This robust system utilizes a serum-free medium that is adaptable for expression of isotopically enriched proteins and carbohydrates for structural studies using mass spectrometry and nuclear magnetic resonance spectroscopy. Furthermore, the composition of the N-glycan can be tuned by adding a small molecule to prevent certain glycan modifications in a manner that does not reduce yield.

Laboratory-scale production of eukaryotic proteins with appropriate post-translational modification represents a significant barrier. Here is a robust protocol with rapid establishment and turnaround for protein expression using a mammalian expression system. This system supports selective amino acid, selective labeling of proteins and small molecule modulators of glycan composition.

The art of producing recombinant proteins with complex post-translational modifications represents a major challenge for studies of structure and ...

Determination of the Relative Cell Surface and Total Expression of Recombinant Ion Channels Using Flow Cytometry

Inherited or de novo mutations in cation-selective channels may lead to sudden cardiac death. Alteration in the plasma membrane trafficking of these multi-spanning transmembrane proteins, with or without change in channel gating, is often postulated to contribute significantly in this process. It has thus become critical to develop a method to quantify the change of the relative cell surface expression of cardiac ion channels on a large scale. Herein, a detailed protocol is provided to determine the relative total and cell surface expression of cardiac L-type calcium channels CaV1.2 and membrane-associated subunits in tsA-201 cells using two-color fluorescent cytometry assays. Compared with other microscopy-based or immunoblotting-based qualitative methods, flow cytometry experiments are fast, reproducible, and large-volume assays that deliver quantifiable end-points on large samples of live cells (ranging from 104 to 106 cells) with similar cellular characteristics in a single flow. Constructs were designed to constitutively express mCherry at the intracellular C-terminus (thus allowing a rapid assessment of the total protein expression) and express an extracellular-facing hemagglutinin (HA) epitope to estimate the cell surface expression of membrane proteins using an anti-HA fluorescence conjugated antibody. To avoid false negative, experiments were also conducted in permeabilized cells to confirm the accessibility and proper expression of the HA epitope. The detailed procedure provides: (1) design of tagged DNA (deoxyribonucleic acid) constructs, (2) lipid-mediated transfection of constructs in tsA-201 cells, (3) culture, harvest, and staining of non-permeabilized and permeabilized cells, and (4) acquisition and analysis of fluorescent signals. Additionally, the basic principles of flow cytometry are explained and the experimental design, including the choice of fluorophores, titration of the HA antibody and control experiments, is thoroughly discussed. This specific approach offers objective relative quantification of the total and cell surface expression of ion channels that can be extended to study ion pumps and plasma membrane transporters.

Inherited cardiac arrhythmias are often caused by mutations that alter the surface delivery of one or more ion channels. Here, we adapt flow cytometry assays to provide a quantification of the relative total and cell surface protein expression of recombinant ion channels expressed in tsA-201 cells.

Inherited or de novo mutations in cation-selective channels may lead to sudden cardiac death. Alteration in the plasma membrane trafficking of these ...