Name: isolation of translating ribosomes containing peptidyl-trnas for functional and structural analyses
Uploaded: 2011-02-25T16:03:00
Duration: 11 min 19 s
Description: Watch this Scientific Journal Video about Isolation of Translating Ribosomes Containing Peptidyl-tRNAs for Functional and Structural Analyses at JoVE.com

L.R.C.-V. wishes to dedicate this paper to the memory of Dr. Adriel D. Johnson, a wonderful Professor whose number one priority was always the education of students. We miss you, Adriel. We are grateful to Jacqueline Moreno for her help in performing the experiments described in this study. This study was supported by a grant provided to C.Y. from the National Science Foundation, MCB-0615390, and by start-up funds provided to L.R.C.-V. from the University of Alabama at Huntsville.

1. Cell Free Extract Preparation.
<ol>
<li>Two grams of a dry Escherichia coli bacterial pellet obtained from mid-log phase cultures are washed, twice, with half liter of buffer A containing 10 mM Tris-acetate, pH 8.0, 14 mM magnesium acetate, 60 mM potassium acetate, and 50 &mu;g/ mL phenylmethanesulphonylfluoride (PMSF) by centrifugation at 5000 X g for 5 min at 4&deg;C. After washing, the bacterial pellet is resuspended in 40 mL of buffer A.</li>
<li>The bacterial cells are disrupted using a French Press, applying 6000 psi pressure. This pressure corresponds to 600 units using a one-inch piston. The disrupted suspension is collected in a clean glass tube (50 mL).</li>
<li>The disrupted suspension is treated with 1 mM dithiotreitol (DTT) and is centrifuged at 30,000 x g for 30 min. The centrifugation procedure is repeated, using the resulting supernatant. The final supernatant is dispensed (1 mL) into microtubes. Finally, the aliquots are frozen using a dry ice-ethanol mixture or liquid nitrogen, and are stored at -60&deg;C or -80&deg;C.</li>
</ol>
2. Preparation of Cell Free Extracts Depleted of RF2.
<ol>
<li>4.5 mL of protein A sepharose 4B slurry beads are mixed with 5 mL of an anti-RF2 anti-serum using a rotating wheel at room temperature, for one hour. The mixture is centrifuged at 2500 x g to separate the beads from the antiserum. Upon discarding the supernatant, these beads containing the anti-RF2 antibodies (anti-RF2 beads) are later washed by resuspension in 1 mL of buffer B containing 35 mM Tris-acetate pH 7.8, 10 mM magnesium acetate, 30 mM ammonium acetate, 60 mM potassium glutamate and 5 &mu;g/ mL of leupeptin proteinase inhibitor. The anti-RF2 beads are then separated from buffer B by centrifugation using the same conditions indicated above. The washing procedure is repeated twice.</li>
<li>One mL of the cell-free extract is mixed with 150 &mu;L of anti-RF2 beads using a rotating wheel, at 4&deg;C, for two hours. The mixture is centrifuged at 10,000 x g to separate the cell-free extract from the beads. The supernatant is removed and treated again with another 150 &mu;L of anti-RF2 beads. This procedure is repeated once more with the resulting cell-free extract solution. The final cell-free extract solution is dispensed (100 &mu;L) into microtubes. The aliquots are frozen using a dry ice-ethanol mixture or liquid nitrogen, and are stored at -60&deg;C or -80&deg;C.</li>
</ol>
3. Preparation of a DNA Template.
<ol>
<li>Prepare a 1 mL PCR-reaction by mixing 600 femtomoles of the plasmid template containing the sequences to be translated (Fig 1), with 0.4 nanomoles of each of the oligodeoxynucleotides indicated in Fig 1, 0.2 micromoles of each dNTP and 50 U of Taq-DNA polymerase in the buffer supplied by Stratagene Co. Perform the amplification reaction under the follow conditions: 94&deg;C for 2 min, (94&deg;C for 30 s, 55 &deg;C for 30 s, 72&deg;C for 1 min; for 30 cycles) and 72&deg;C for 12 min.</li>
<li>Purify the PCR-products by precipitating the DNA by adding 1/10 volume of 3 M sodium acetate, pH 5.2, and 2 volumes of ice-cold ethanol. Repeat the precipitation procedure once more. Resuspend the DNA in 100 &mu;L of diethyl-pyrocarbonate (DEPC) treated-water. 
Typically, this procedure yields 100 &mu;g of a 600 bp DNA product.</li>
<li>Verify the integrity of the PCR products by electrophoresis on agarose gels.</li>
</ol>
4. Preparation of Biotin Labeled mRNA.
<ol>
<li>Prepare a 100 &mu;L in vitro transcription reaction in DEPC treated water by mixing 5 &mu;g of the PCR-generated DNA fragment with 0.5 micromoles of ATP, CTP, GTP, 0.3 micromoles of UTP, 50 nanomoles of biotin-16-UTP, and 10 &mu;L of T7 enzyme mix in the buffer supplied by Promega Co. Incubate the reaction mixture at 37&deg;C for 3 hours. 
To quantify the amount of mRNA obtained, the DNA template should be eliminated by adding RNase-free DNase. Incubate the reaction at 37&deg;C for 10 min.</li>
<li>Purify the mRNA products by precipitating the RNA by adding 1/10 volume of 3 M sodium acetate, pH 5.2, and 2 volumes of cold ethanol. Repeat the precipitation procedure once more. Resuspend the mRNA in 100 &mu;L of DEPC treated-water. 
Usually, this procedure yields between 1-2 mg of a 600 nt biotin labeled mRNA product.</li>
<li>Verify the integrity of the biotin labeled mRNA products by electrophoresis on agarose gels.</li>
</ol>
5. Isolation of Translating Ribosomes Containing a Peptidyl-tRNA.
<ol>
<li>Prepare 500 &mu;L of an in vitro translation reaction mixture in DEPC treated water by mixing 10-15 &mu;g of biotin-labeled mRNA with 75 nanomoles of each of the nineteen amino acids -all except the amino acid that will be replaced by a radioactive amino acid, 50 &mu;Ci of a radioactive amino acid (37MBq), and 20-50 &mu;L of RF2-depleted cell-free extract, in a buffered reaction mixture containing 40 mM Tris-acetate, pH 8.0, 10 mM magnesium acetate, 175 mM potassium acetate, 10 mM ammonium acetate, 2 mM DTT, 2 mM ATP, 0.5 mM GTP, 30 mM phosphoenolpyruvate (PEP), 0.3 U/ mL pyruvate kinase (PK), 3.5% polyethylene glycol 8000, 1 mM spermidine, 20 &mu;g/ mL folinic acid, and 250 &mu;g/ mL tRNA from E. coli. Incubate the reaction mixture at 37&deg;C for 10 minutes. 
The order of addition of the reaction components is very important. The cell-free extract must be mixed first with the buffer solution containing the amino acids and the final mixture incubated for 5 min at room temperature to allow the activation of the ribosomes. Later, the biotin-labeled mRNAs are added. For structural analyses using chemical modification procedures, the addition of a radioactive amino acid is not required.</li>
<li>Add to the translation reaction mixture - 3 mL of streptavidin paramagnetic beads (SMB) suspended in buffer C containing 35 mM Tris-acetate, pH 8.0, 10 mM magnesium acetate, 175 mM potassium acetate, 10 mM ammonium acetate and 1 mM DTT. Incubate the new suspension at room temperature for 10 min.</li>
<li>Separate the SMB from the mixture by applying a magnetic field (MF) using magnetic separation stands.</li>
<li>Resuspend the SMB in buffer C and separate the beads again using the MF. Repeat this washing procedure twice. Resuspend the beads in 500 &mu;L of buffer C, store the suspension on ice, and perform the next procedure immediately.</li>
</ol>
6. Analysis of the Isolated Ribosomes Containing the Peptidyl-tRNA.

For peptidyl-tRNA
<ol>
<li>Mix 10 &mu;L of the SMB (suspended in buffer C) with 10 &mu;L of a loading buffer containing 50 mM Tris-HCl, pH 6.8, 2.5% (v/v) glycerol, 4% sodium dodecyl sulfate, 2 mM DTT and 0.2 mg/ mL bromophenol blue. Resolve the components attached to the beads by running the samples in 10% Tris-tricine polyacrylamide gels.</li>
<li>Dry the gels using a vacuum gel-dryer.</li>
<li>Verify the integrity and purification of the peptidyl-tRNA by exposing the dried gel to an X-ray film.</li>
</ol>
For ribosomal RNA
<ol start="4">
<li>Add 190 &mu;L of 2 mM ethylenediaminetetraacetate (EDTA) solution prepared with DEPC-treated water, and 200 &mu;L of phenol equilibrated with water, to 10 &mu;L of a bead suspension. Mix the suspension by vigorous vortexing and separate the inorganic phase from the organic phase by centrifugation at 10,000 x g for 3 min at room temperature. Collect the top water layer in a new microtube.</li>
<li>Precipitate the RNA from the water layer by adding 1/10 volume of 3 M sodium acetate, pH 5.2, 1 &mu;L of 20 mg/ mL of glycogen solution, and 2 volumes of ice-cold ethanol. Resuspend the RNA in 10 &mu;L of DEPC-treated water.</li>
<li>Verify the integrity of the ribosomal RNAs by electrophoresis on agarose gels. 
Usually, 50 &mu;L of beads suspension yields 1&mu;g of ribosomal RNA.</li>
</ol>
7. Representative Results:

Figure 2 presents the results of a series of analyses evaluating the quality and functionality of translating ribosomes isolated using the procedure outlined. The observation of a unique band resolved in polyacrylamide gels indicates the presence of polypeptides bound to the biotin-labeled mRNAs attached to the SMB (Figure 2A). The purification of ribosomal RNAs using this procedure establishes the presence of ribosomes bound to these biotin-labeled mRNAs, as well (Figure 2B). Addition of puromycin, an antibiotic that induces the peptidyl-transferase activity of the ribosome, results in cleavage of the nascent peptidyl-tRNAs1. This is observed as a shift in the migration pattern of the isolated polypeptide in polyacrylamide gels (Fig 2C). Together, these data indicate that biotin-labeled mRNAs attached to the SMB contain functional ribosomes with peptidyl-tRNAs.
The isolation of translating ribosomes containing specific peptidyl-tRNAs permits the study of the effects of peptidyl-tRNAs, antibiotics, and other molecules, on ribosome function (Figure 3A), and on ribosome structure (Figure 3B). In the examples presented here the antibiotic sparsomycin and the amino acid tryptophan (Trp) inhibit the hydrolytic cleavage of the nascent tnaC-tRNA peptidyl-tRNA induced by RF2 in the isolated ribosomes (Figure 3A). The interaction of sparsomycin or Trp with the ribosome also induces structural changes in some nucleotides that constitute the 23S rRNA of the translating ribosome (Fig 3B). In summary, the biological material obtained using the procedure described here are useful in obtaining additional understanding of the structural contacts involved in inhibition of ribosome function upon its interaction with various molecular factors.

 <img src="/files/ftp_upload/2498/2498fig1.jpg" alt="Figure 1" /> 
Figure 1. Procedure used to produce and purify translating ribosomes containing peptidyl-tRNAs. DNA fragments ca. 650 bp in length, containing the tnaC gene and its adjacent region, shown in the figure, are used as templates in preparing mRNAs containing biotin, [(bio)UMP]mRNA. These DNA fragments are produced by PCR procedures using the oligodeoxynucleotides indicated at the top of the figure. Underlined letters mark the location of the T7 promoter encoded in the nucleotide primer. Bold letters indicate the Shine-Dalgarno sequence that is used in the translation of the tnaC mRNA sequences. The T7 promoter in the DNA fragment is recognized by T7 RNA polymerase which is used to transcribe (arrow) the tnaC sequence and the non-coding sequence downstream. Transcription is terminated when RNA polymerase reaches the 3'-end of the DNA fragment, after the rpoBC terminator sequences (rpoBC"t"). The stem-loop structure of the rpoBC terminator protects the mRNA from the 3'-5' exoribonuclease activity present in the cell-free extracts used in the in vitro translation reactions. The isolated [(bio)UMP]mRNAs are used to generate stalled, translating ribosomes, using in vitro translation assays. The tnaC sequence in the [(bio)UMP]mRNA is translated by a ribosome which stalls when the last amino acid is incorporated, due to the absence of the Release Factor (RF2) from the reaction mixture. The ribosome-[(bio)UMP]mRNA complexes are isolated from the total reaction mixture using streptavidin-magnetic-beads (SMB) which bind the biotin-labeled nucleotides incorporated in the mRNA. The SMBs bound to the ribosome-[(bio)UMP]mRNA complexes are extracted from the total reaction mixture by using a magnetic field (MF).

 <img src="/files/ftp_upload/2498/2498fig2.jpg" alt="Figure 2" /> 
Figure 2. Structural and functional analyses of the isolated complexes. A) In vitro translation reactions were performed with [(bio)UMP]mRNAs in which the start codon of the tnaC gene was replaced by a stop codon. The final products were isolated using streptavidin beads, as is indicated in Fig 1. Reaction products and isolated molecules were then analyzed by electrophoresis on polyacrylamide gels. The tnaC-tRNA and tnaC peptide bands are marked. B) Total RNA was isolated from the stalled complexes using phenol extraction procedures. Each isolated RNA was resolved by electrophoresis in agarose gels. Ribosomal RNAs (rRNAs) are indicated. C) Isolated complexes containing tnaC-tRNAs were incubated with (+) or without (-) 0.02 mM puromycin (Puro) at room temperature for 10 min. The tnaC peptide were labeled during translation using [35S]-methionine (A) or [3H]-proline (B).

 <img src="/files/ftp_upload/2498/2498fig3.jpg" alt="Figure 3" /> 
Figure 3. Structural and functional analyses of ribosomes containing tnaC-tRNA that are bound to the antibiotic sparsomycin, or the amino acid tryptophan. A) Isolated complexes containing tnaC-tRNAs were incubated with (+) or without (-) sparsomycin (Spar) or tryptophan (Trp) at room temperature for 5 min. Later, the mixtures were mixed with RF2 and incubated at 37&deg;C for 20 min. B) Analysis of methylation changes in 23S rRNA induced by molecules binding within the ribosome. Isolated complexes were pre-incubated with (+) or without (-) 2 mM Trp or Spar for 5 min at 37&deg;C. Then, an alkylation agent, dimethyl sulfate, was added and the mixtures incubated at room temperature for an additional 20 min. Total RNA was extracted and primer extension assays were performed with a [32P]-labeled oligodeoxynucleotide complementary to the nucleotides of 23S rRNA which are spaced 100 nucleotides downstream from the modified nucleotide. The final products of the extension assays were resolved by electrophoresis in polyacrylamide gels. The 23S rRNA nucleotides affected by the presence of Trp or Spar are indicated.

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No conflicts of interest declared.

This isolation procedure described in this report is highly reproducible. Unfortunately, the presence of short peptidyl-tRNAs produced from the same translated sequences cannot be conveniently avoided; such peptidyl-tRNAs may correspond to 15-20 % of the total isolated material (Figure 2C). These contaminants may increase in concentration when higher concentrations of biotin-labeled mRNA have been used or when the cell-free extracts used are old or have been re-used several times. Therefore, it is very important to control the quality and concentration of the components of the in vitro translation reactions. Alternatively, commercial high efficiency reconstituted cell-free extracts are available that can be used for the same purposes (PURE system)18.
Using this method, biotin-16-UMP is randomly incorporated along the length of the target mRNA. There is a possibility that some ORFs containing uridine tracts may have incorporated this modified nucleotide in their sequences, affecting their translation. Variable relative concentrations of biotin-16-UTP/UTP have to be tested during synthesis of the mRNA in order to obtain the most efficient translation. Alternative methods of biotin labeling can be used, targeting the 5'-end, the most stable end of the mRNAs. (i) Biotin can be attached to the 5'-end of the mRNA using 3'-NH2ATP (3'-amino, 3'-deoxyadenosine triphosphate) and T4 RNA ligase19. (ii) 5'-end biotin-labeled oligonucleotide can be attached to the 5'-end of the mRNA using T4 RNA ligase as well20. (iii) 3'-end biotin-labeled oligodeoxynucleotides which correspond to the complementary sequence at the 5'-end of the mRNA could also be used to isolate the translating complexes. These biotin labeled oligodeoxynucleotides could be attached to the SMB prior to isolation. Later the SMB attached to the biotin-labeled oligodeoxynucleotides could be mixed with the translation reaction mixture to allow interaction with their complementary mRNA sequences. After isolation, the hybrid RNA-DNA region - and the region hybridizing between the biotin-labeled oligodeoxynucleotide and the mRNA sequence - could be degraded using RNAse H, separating the stalled complexes from the streptavidin beads. This alternative method may allow the complexes to be employed in future structural analyses using higher resolution methods.

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		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
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		<tbody>
		
<tr>
<td>Tris base</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>252859</td>
<td>&nbsp;</td>
<tr>
<td>Magnesium acetate</td>
<td>&nbsp;</td>
<td>Fluka</td>
<td>63049</td>
<td>&nbsp;</td>
<tr>
<td>Potassium glutamate</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>G1501</td>
<td>&nbsp;</td>
<tr>
<td>Ammonium acetate</td>
<td>&nbsp;</td>
<td>Fluka</td>
<td>09688</td>
<td>&nbsp;</td>
<tr>
<td>PMSF</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P7626</td>
<td>&nbsp;</td>
<tr>
<td>Protein A sepharose 4B slurry beads</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P9424</td>
<td>&nbsp;</td>
<tr>
<td>Leupeptin inhibitor</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>L9783</td>
<td>&nbsp;</td>
<tr>
<td>Taq-DNA polymerase</td>
<td>&nbsp;</td>
<td>Stratagene</td>
<td>201223</td>
<td>&nbsp;</td>
<tr>
<td>T7 Maxiprep kit</td>
<td>&nbsp;</td>
<td>Promega</td>
<td>P1300</td>
<td>&nbsp;</td>
<tr>
<td>SMB</td>
<td>&nbsp;</td>
<td>Promega</td>
<td>Z5482</td>
<td>&nbsp;</td>
<tr>
<td>Biotin-16-UTP</td>
<td>&nbsp;</td>
<td>Roche</td>
<td>1388908</td>
<td>&nbsp;</td>
<tr>
<td>ATP</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>A7699</td>
<td>&nbsp;</td>
<tr>
<td>GTP</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>G8877</td>
<td>&nbsp;</td>
<tr>
<td>PEP</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P7002</td>
<td>&nbsp;</td>
<tr>
<td>PK</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P7768</td>
<td>&nbsp;</td>
<tr>
<td>Polyethylenglycol 8000</td>
<td>&nbsp;</td>
<td>Fluka</td>
<td>81268</td>
<td>&nbsp;</td>
<tr>
<td>Folinic acid</td>
<td>&nbsp;</td>
<td>Fluka</td>
<td>47612</td>
<td>&nbsp;</td>
<tr>
<td>Spermidine</td>
<td>&nbsp;</td>
<td>Fluka</td>
<td>85558</td>
<td>&nbsp;</td>
<tr>
<td>tRNA from E. coli</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>R1753</td>
<td>&nbsp;</td>
<tr>
<td>DEPC</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>D5758</td>
<td>&nbsp;</td>
<tr>
<td>Tricine</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>T5816</td>
<td>&nbsp;</td>
<tr>
<td>L-amino acids</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>LAA21</td>
<td>&nbsp;</td>
<tr>
<td>DTT</td>
<td>&nbsp;</td>
<td>Fluka</td>
<td>43815</td>
<td>&nbsp;</td>
<tr>
<td>magnetic separation stands</td>
<td>&nbsp;</td>
<td>Promega</td>
<td>Z5342</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

isolation of translating ribosomes containing peptidyl-trnas for functional and structural analyses

Recently, structural and biochemical studies have detailed many of the molecular events that occur in the ribosome during inhibition of protein synthesis by antibiotics and during nascent polypeptide synthesis. Some of these antibiotics, and regulatory nascent polypeptides mostly in the form of peptidyl-tRNAs, inhibit either peptide bond formation or translation termination1-7. These inhibitory events can stop the movement of the ribosome, a phenomenon termed "translational arrest". Translation arrest induced by either an antibiotic or a nascent polypeptide has been shown to regulate the expression of genes involved in diverse cellular functions such as cell growth, antibiotic resistance, protein translocation and cell metabolism8-13. Knowledge of how antibiotics and regulatory nascent polypeptides alter ribosome function is essential if we are to understand the complete role of the ribosome in translation, in every organism.
Here, we describe a simple methodology that can be used to purify, exclusively, for analysis, those ribosomes translating a specific mRNA and containing a specific peptidyl-tRNA14. This procedure is based on selective isolation of translating ribosomes bound to a biotin-labeled mRNA. These translational complexes are separated from other ribosomes in the same mixture, using streptavidin paramagnetic beads (SMB) and a magnetic field (MF). Biotin-labeled mRNAs are synthesized by run-off transcription assays using as templates PCR-generated DNA fragments that contain T7 transcriptional promoters. T7 RNA polymerase incorporates biotin-16-UMP from biotin-UTP; under our conditions approximately ten biotin-16-UMP molecules are incorporated in a 600 nt mRNA with a 25% UMP content. These biotin-labeled mRNAs are then isolated, and used in in vitro translation assays performed with release factor 2 (RF2)-depleted cell-free extracts obtained from Escherichia coli strains containing wild type or mutant ribosomes. Ribosomes translating the biotin-labeled mRNA sequences are stalled at the stop codon region, due to the absence of the RF2 protein, which normally accomplishes translation termination. Stalled ribosomes containing the newly synthesized peptidyl-tRNA are isolated and removed from the translation reactions using SMB and an MF. These beads only bind biotin-containing messages.
The isolated, translational complexes, can be used to analyze the structural and functional features of wild type or mutant ribosomal components, or peptidyl-tRNA sequences, as well as determining ribosome interaction with antibiotics or other molecular factors 1,14-16. To examine the function of these isolated ribosome complexes, peptidyl-transferase assays can be performed in the presence of the antibiotic puromycin1. To study structural changes in translational complexes, well established procedures can be used, such as i) crosslinking to specific amino acids14 and/or ii) alkylation protection assays1,14,17.

Watch this Scientific Journal Video about Isolation of Translating Ribosomes Containing Peptidyl-tRNAs for Functional and Structural Analyses at JoVE.com

Isolation of Translating Ribosomes Containing Peptidyl-tRNAs for Functional and Structural Analyses

A major impediment to biochemical analyses of ribosomes containing nascent peptidyl-tRNAs has been the presence of other ribosomes in the same samples, ribosomes not involved in the translation of the specific mRNA sequence being analyzed. We developed a simple methodology to purify, exclusively, the ribosomes containing the nascent peptidyl-tRNA of interest.

Recently, structural and biochemical studies have detailed many of the molecular events that occur in the ribosome during inhibition of protein synthesis ...

isolation-translating-ribosomes-containing-peptidyl-trnas-for

Research

JoVE Journal

Biology

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

Evisceration of Mouse Vitreous and Retina for Proteomic Analyses

While the mouse retina has emerged as an important genetic model for inherited retinal disease, the mouse vitreous remains to be explored. The vitreous is a highly aqueous extracellular matrix overlying the retina where intraocular as well as extraocular proteins accumulate during disease.1-3 Abnormal interactions between vitreous and retina underlie several diseases such as retinal detachment, proliferative diabetic retinopathy, uveitis, and proliferative vitreoretinopathy.1,4 The relative mouse vitreous volume is significantly smaller than the human vitreous (Figure 1), since the mouse lens occupies nearly 75% of its eye.5 This has made biochemical studies of mouse vitreous challenging. In this video article, we present a technique to dissect and isolate the mouse vitreous from the retina, which will allow use of transgenic mouse models to more clearly define the role of this extracellular matrix in the development of vitreoretinal diseases.

The dissection technique illustrates evisceration of the vitreous, retina, and lens from the mouse eye, separation by centrifugation, and characterization with protein assays.

While the mouse retina has emerged as an important genetic model for inherited retinal disease, the mouse vitreous remains to be explored. The vitreous is ...

Isolation and Characterization of RNA-Containing Exosomes

The field of exosome research is rapidly expanding, with a dramatic increase in publications in recent years. These small vesicles (30-100 nm) of endocytic origin were first proposed to function as a way for reticulocytes to eradicate the transferrin receptor while maturing into erythrocytes1, and were later named exosomes. Exosomes are formed by inward budding of late endosomes, producing multivesicular bodies (MVBs), and are released into the environment by fusion of the MVBs with the plasma membrane2. Since the first discovery of exosomes, a wide range of cells have been shown to release these vesicles. Exosomes have also been detected in several biological fluids, including plasma, nasal lavage fluid, saliva and breast milk3-6. Furthermore, it has been demonstrated that the content and function of exosomes depends on the originating cell and the conditions under which they are produced. A variety of functions have been demonstrated for exosomes, such as induction of tolerance against allergen7,8, eradication of established tumors in mice9, inhibition and activation of natural killer cells10-12, promotion of differentiation into T regulatory cells13, stimulation of T cell proliferation14 and induction of T cell apoptosis15. Year 2007 we demonstrated that exosomes released from mast cells contain messenger RNA (mRNA) and microRNA (miRNA), and that the RNA can be shuttled from one cell to another via exosomes. In the recipient cells, the mRNA shuttled by exosomes was shown to be translated into protein, suggesting a regulatory function of the transferred RNA16. Further, we have also shown that exosomes derived from cells grown under oxidative stress can induce tolerance against further stress in recipient cells and thus suggest a biological function of the exosomal shuttle RNA17. Cell culture media and biological fluids contain a mixture of vesicles and shed fragments. A high quality isolation method for exosomes, followed by characterization and identification of the exosomes and their content, is therefore crucial to distinguish exosomes from other vesicles and particles. Here, we present a method for the isolation of exosomes from both cell culture medium and body fluids. This isolation method is based on repeated centrifugation and filtration steps, followed by a final ultracentrifugation step in which the exosomes are pelleted. Important methods to identify the exosomes and characterize the exosomal morphology and protein content are highlighted, including electron microscopy, flow cytometry and Western blot. The purification of the total exosomal RNA is based on spin column chromatography and the exosomal RNA yield and size distribution is analyzed using a Bioanalyzer.

This paper demonstrates methods for the isolation, purification and detection of exosomes, as well as techniques for analysis of their molecular content. These methods are adaptable for exosome isolation from both cell culture media and biological fluids, and can beyond analysis of molecular content also be useful in functional studies.

The field of exosome research is rapidly expanding, with a dramatic increase in publications in recent years. These small vesicles (30-100 nm) of ...

Chromatographic Purification of Highly Active Yeast Ribosomes

Eukaryotic ribosomes are much more labile as compared to their eubacterial and archael counterparts, thus posing a significant challenge to researchers. Particularly troublesome is the fact that lysis of cells releases a large number of proteases and nucleases which can degrade ribosomes. Thus, it is important to separate ribosomes from these enzymes as quickly as possible. Unfortunately, conventional differential ultracentrifugation methods leaves ribosomes exposed to these enzymes for unacceptably long periods of time, impacting their structural integrity and functionality. To address this problem, we utilize a chromatographic method using a cysteine charged Sulfolink resin. This simple and rapid application significantly reduces co-purifying proteolytic and nucleolytic activities, producing high yields of intact, highly biochemically active yeast ribosomes. We suggest that this method should also be applicable to mammalian ribosomes. The simplicity of the method, and the enhanced purity and activity of chromatographically purified ribosome represents a significant technical advancement for the study of eukaryotic ribosomes.

Contamination of preparations of eukaryotic ribosomes purified by traditional methods by co-purifying nucleases and proteases negatively impacts on downstream biochemical and structural analyses. A rapid and simple chromatographic purification method is used to solve this problem using yeast ribosomes as a model system.

Eukaryotic ribosomes are much more labile as compared to their eubacterial and archael counterparts, thus posing a significant challenge to researchers.  ...

Use of a Robot for High-throughput Crystallization of Membrane Proteins in Lipidic Mesophases

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through macromolecular X-ray crystallography (MX). An essential ingredient of MX is a steady supply of ideally diffraction-quality crystals. The in meso or lipidic cubic phase (LCP) method for crystallizing membrane proteins is one of several methods available for crystallizing membrane proteins. It makes use of a bicontinuous mesophase in which to grow crystals. As a method, it has had some spectacular successes of late and has attracted much attention with many research groups now interested in using it. One of the challenges associated with the method is that the hosting mesophase is extremely viscous and sticky, reminiscent of a thick toothpaste. Thus, dispensing it manually in a reproducible manner in small volumes into crystallization wells requires skill, patience and a steady hand. A protocol for doing just that was developed in the Membrane Structural &amp; Functional Biology (MS&amp;FB) Group1-3. JoVE video articles describing the method are available1,4. 
The manual approach for setting up in meso trials has distinct advantages with specialty applications, such as crystal optimization and derivatization. It does however suffer from being a low throughput method. Here, we demonstrate a protocol for performing in meso crystallization trials robotically. A robot offers the advantages of speed, accuracy, precision, miniaturization and being able to work continuously for extended periods under what could be regarded as hostile conditions such as in the dark, in a reducing atmosphere or at low or high temperatures. An in meso robot, when used properly, can greatly improve the productivity of membrane protein structure and function research by facilitating crystallization which is one of the slow steps in the overall structure determination pipeline. 
In this video article, we demonstrate the use of three commercially available robots that can dispense the viscous and sticky mesophase integral to in meso crystallogenesis. The first robot was developed in the MS&amp;FB Group5,6. The other two have recently become available and are included here for completeness.	
An overview of the protocol covered in this article is presented in Figure 1. All manipulations were performed at room temperature (~20 &deg;C) under ambient conditions.

Herein is described a robotic approach to high-throughput crystallization of membrane proteins in lipidic mesophases for use in structure determination using macromolecular X-ray crystallography. Three robots capable of handling the viscous and sticky protein-laden mesophase integral to the method are introduced.

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through ...

Harvesting and Cryo-cooling Crystals of Membrane Proteins Grown in Lipidic Mesophases for Structure Determination by Macromolecular Crystallography

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at atomic resolution. Presently, there is only one way to capture such information as applied to integral membrane proteins (Figure 1), and the complexes they form, and that method is macromolecular X-ray crystallography (MX). To do MX diffraction quality crystals are needed which, in the case of membrane proteins, do not form readily. A method for crystallizing membrane proteins that involves the use of lipidic mesophases, specifically the cubic and sponge phases1-5, has gained considerable attention of late due to the successes it has had in the G protein-coupled receptor field6-21 (<a href="http://www.mpdb.tcd.ie" target="_blank">www.mpdb.tcd.ie</a>). However, the method, henceforth referred to as the in meso or lipidic cubic phase method, comes with its own technical challenges. These arise, in part, due to the generally viscous and sticky nature of the lipidic mesophase in which the crystals, which are often micro-crystals, grow. Manipulating crystals becomes difficult as a result and particularly so during harvesting22,23. Problems arise too at the step that precedes harvesting which requires that the glass sandwich plates in which the crystals grow (Figure 2)24,25 are opened to expose the mesophase bolus, and the crystals therein, for harvesting, cryo-cooling and eventual X-ray diffraction data collection.
The cubic and sponge mesophase variants (Figure 3) from which crystals must be harvested have profoundly different rheologies4,26. The cubic phase is viscous and sticky akin to a thick toothpaste. By contrast, the sponge phase is more fluid with a distinct tendency to flow. Accordingly, different approaches for opening crystallization wells containing crystals growing in the cubic and the sponge phase are called for as indeed different methods are required for harvesting crystals from the two mesophase types. Protocols for doing just that have been refined and implemented in the Membrane Structural and Functional Biology (MS&amp;FB) Group, and are described in detail in this JoVE article (Figure 4). Examples are given of situations where crystals are successfully harvested and cryo-cooled. We also provide examples of cases where problems arise that lead to the irretrievable loss of crystals and describe how these problems can be avoided. In this article the Viewer is provided with step-by-step instructions for opening glass sandwich crystallization wells, for harvesting and for cryo-cooling crystals of membrane proteins growing in cubic and in sponge phases.

Herein is described procedures implemented in the Caffrey Membrane Structural and Functional Biology Group to harvest and cryo-cool membrane protein crystals grown in lipidic cubic and sponge phases for use in structure determination using macromolecular X-ray crystallography.

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at ...

Isolation, Culture, and Functional Characterization of Adult Mouse Cardiomyoctyes

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of heart disease. In particular, the ability to study ion homeostasis, ion channel function, cellular excitability and excitation-contraction coupling and their alterations in diseased conditions and by disease-causing mutations have led to significant insights into cardiac diseases. Furthermore, the lack of an adequate immortalized cell line to mimic adult CMs, and the limitations of neonatal CMs (which lack many of the structural and functional biomechanics characteristic of adult CMs) in culture have hampered our understanding of the complex interplay between signaling pathways, ion channels and contractile properties in the adult heart strengthening the importance of studying adult isolated cardiomyocytes. Here, we present methods for the isolation, culture, manipulation of gene expression by adenoviral-expressed proteins, and subsequent functional analysis of cardiomyocytes from the adult mouse. The use of these techniques will help to develop mechanistic insight into signaling pathways that regulate cellular excitability, Ca2+ dynamics and contractility and provide a much more physiologically relevant characterization of cardiovascular disease.

Here we describe the isolation of adult mouse cardiomyoctyes using a Langendorff perfusion system. The resulting cells are Ca2+-tolerant, electrically quiescent and can be cultured and transfected with adeno- or lentiviruses to manipulate gene expression. Their functionality can also be analyzed using the MMSYS system and patch clamp techniques.

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of ...

Reconstitution of a Kv Channel into Lipid Membranes for Structural and Functional Studies

To study the lipid-protein interaction in a reductionistic fashion, it is necessary to incorporate the membrane proteins into membranes of well-defined lipid composition. We are studying the lipid-dependent gating effects in a prototype voltage-gated potassium (Kv) channel, and have worked out detailed procedures to reconstitute the channels into different membrane systems. Our reconstitution procedures take consideration of both detergent-induced fusion of vesicles and the fusion of protein/detergent micelles with the lipid/detergent mixed micelles as well as the importance of reaching an equilibrium distribution of lipids among the protein/detergent/lipid and the detergent/lipid mixed micelles. Our data suggested that the insertion of the channels in the lipid vesicles is relatively random in orientations, and the reconstitution efficiency is so high that no detectable protein aggregates were seen in fractionation experiments. We have utilized the reconstituted channels to determine the conformational states of the channels in different lipids, record electrical activities of a small number of channels incorporated in planar lipid bilayers, screen for conformation-specific ligands from a phage-displayed peptide library, and support the growth of 2D crystals of the channels in membranes. The reconstitution procedures described here may be adapted for studying other membrane proteins in lipid bilayers, especially for the investigation of the lipid effects on the eukaryotic voltage-gated ion channels.

Procedures for complete reconstitution of a prototype voltage-gated potassium channel into lipid membranes are described. The reconstituted channels are suitable for biochemical assays, electrical recordings, ligand screening and electron crystallographic studies. These methods may have general applications to the structural and functional studies of other membrane proteins.

To study the lipid-protein interaction in a reductionistic fashion, it is necessary to incorporate the membrane proteins into membranes of well-defined ...

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of cell heath due to their role in both cell metabolism and energy production as well as control of apoptosis. Therefore, direct evaluation of isolated mitochondria and mitochondrial perturbation offers the ability to determine if organelle-specific (dys)function is occurring. The methods described in this protocol include isolation of intact, functional mitochondria from HEK cultured cells and mouse liver and spinal cord, but can be easily adapted for use with other cultured cells or animal tissues. Mitochondrial function assessed by TMRE and the use of common mitochondrial uncouplers and inhibitors in conjunction with a fluorescent plate reader allow this protocol not only to be versatile and accessible to most research laboratories, but also offers high throughput.

Comparison of mitochondrial membrane potential between samples yields valuable information about cellular status. Detailed steps for isolating mitochondria and assessing response to inhibitors and uncouplers using fluorescence are described. The method and utility of this protocol are illustrated by use of a cell culture and animal model of cellular stress.

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of ...

A Rapid, Scalable Method for the Isolation, Functional Study, and Analysis of Cell-derived Extracellular Matrix

The extracellular matrix (ECM) is recognized as a diverse, dynamic, and complex environment that is involved in multiple cell-physiological and pathological processes. However, the isolation of ECM, from tissues or cell culture, is complicated by the insoluble and cross-linked nature of the assembled ECM and by the potential contamination of ECM extracts with cell surface and intracellular proteins. Here, we describe a method for use with cultured cells that is rapid and reliably removes cells to isolate a cell-derived ECM for downstream experimentation.
Through use of this method, the isolated ECM and its components can be visualized by in situ immunofluorescence microscopy. The dynamics of specific ECM proteins can be tracked by tracing the deposition of a tagged protein using fluorescence microscopy, both before and after the removal of cells. Alternatively, the isolated ECM can be extracted for biochemical analysis, such as sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblotting. At larger scales, a full proteomics analysis of the isolated ECM by mass spectrometry can be conducted. By conducting ECM isolation under sterile conditions, sterile ECM layers can be obtained for functional or phenotypic studies with any cell of interest. The method can be applied to any adherent cell type, is relatively easy to perform, and can be linked to a wide repertoire of experimental designs.

The extracellular matrix plays a major role in defining the microenvironment of cells and in modulating cell behavior and phenotype. We describe a rapid method for the isolation of cell-derived extracellular matrix, which can be adapted to different scales for microscopic, biochemical, proteomic, or functional studies.