This work was supported by the University of Western Ontario, the Department of Obstetrics and Gynaecology; and a grant ER06-02-188 from the Ministryof Research and Innovation, Early Researcher Award. MMD was supported by a CIHR Training Program in Reproduction, Early Development and the Impact on Health (REDIH) Graduate Scholarship.

DAY 1
Prepare the following solutions fresh on the day of oocyte collection with sterile, distilled water such as GIBCO water. To reduce the chance of DNA contamination, change gloves often and use filter tips. Keep tubes angled away when open, and recap all tubes when not in use. We recommend that solutions are made as n+1.
3% LMP Agarose
30 mg Low Melting Point (LMP) Agarose 
up to 1 ml GIBCO H2O 
dissolve @ 70 &deg;C
Lysis Solution
8 &mu;l lysis buffer 
1 &mu;l proteinase K 
1 &mu;l 10% IGEPAL 
place on ice until ready for use
2:1 Agarose:Lysis Solution (10 &mu;l per individual oocyte, amount is for 3 oocytes)
20 &mu;l 3% LMP agarose 
10 &mu;l Lysis Solution 
mix @ 70 &deg;C
SDS Lysis Buffer (501 &mu;l per individual oocyte)
<table border="0">
	<tbody>
		<tr>
			<td>1x TE pH 7.5</td>
			<td>450 &mu;l</td>
		</tr>
		<tr>
			<td>10% SDS</td>
			<td>50 &mu;l</td>
		</tr>
		<tr>
			<td>Proteinase K</td>
			<td>1 &mu;l</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>501 &mu;l</td>
		</tr>
	</tbody>
</table>
1. Oocyte Collection
<ol>
	<li>Place the dissected mouse oviducts in M2 media, and tear the ampullae to extract the cumulus cell complex.</li>
	<li>Separate the oocytes from the cumulus cell complex using 0.3 mg/ml hyaluronidase solution in a 30 &mu;l drop of M2 media. Keep the oocytes in solution only as long as it takes to remove the cumulus cells, as lengthy exposure may damage them. Wash the oocytes 3x in 30 &mu;l drop of M2 media, removing cumulus cells periodically.</li>
	<li>Remove the zona pellucida using acidic tyrode's solution. Place the oocytes in one 30 &mu;l drop of solution first, and then transfer to another 30 &mu;l drop, as any media carried along will dilute the acid and reduce its efficiency. Keep the oocytes in solution only as long as it takes to remove the zona, as lengthy exposure may damage them. Note: an increased concentration of acidic tyrode's solution or pronase may be used for human samples, as the human zona pellucida is more resistant to treatment with acidic tyrode's solution than the mouse.</li>
	<li>Wash the oocytes once more in a 30 &mu;l drop of M2 media.</li>
</ol>
2. Agarose Embedding and Lysis
<ol>
	<li>To perform agarose embedding, place the lysis solution on a 70 &deg;C heatblock. Add the preheated LMP agarose to the lysis solution, producing a 2:1 agarose:lysis solution.</li>
	<li>Place a single oocyte onto a clean glass slide in minimal M2 media. Take up 10 &mu;l of the agarose:lysis solution into a pipette tip, and (under a microscope) gently expel a small amount (~1 &mu;l or less) onto the glass slide, allowing it to mix with the minimal media. Gently pick up the oocyte into the pipette tip and put all 10 &mu;l into an Eppendorf tube with 300 &mu;l mineral oil so the bead forms a sphere. 
	Note: this process must be done fairly quickly as the agarose will harden if the temperature drops as little as 5 &deg;C below 70 &deg;C.</li>
	<li>Incubate the tube on ice for 10 minutes. To perform lysis, remove the 300 &mu;l mineral oil and add 500 &mu;l of the SDS lysis buffer. Incubate overnight in a 50 &deg;C water bath. 
	Note: Lysis solution (Table 1) may also be used for this purpose.</li>
</ol>
DAY 2
Prepare the following solutions fresh on the day of bisulfite mutagenesis. To reduce chance of DNA contamination, change gloves often and use filter tips. Keep tubes angled away when open, and recap all tubes when not in use. We recommend that solutions are made as n+1.
<table border="0">
	<tbody>
		<tr>
			<td>3 M NaOH</td>
			<td>2.4 g NaOH in 20 ml autoclaved ddH2O</td>
		</tr>
		<tr>
			<td>0.1 M NaOH</td>
			<td>0.5 ml of 3M in 14.5 ml autoclaved ddH2O</td>
		</tr>
		<tr>
			<td>0.3 M NaOH</td>
			<td>1.5 ml of 3M in 13.5 ml autoclaved ddH2O</td>
		</tr>
	</tbody>
</table>
2.5 M Bisulfite Solution
<ol class="lalpha">
	<li>3.8 g sodium bisulfite 
	5.5 ml GIBCO distilled H2O 
	1 ml 3 M NaOH 
	dissolve @ room temperature</li>
	<li>110 mg Hydroquinone 
	1 ml GIBCO distilled H2O 
	dissolve @ 90 &deg;C (for only as long as it takes to dissolve, mix regularly)</li>
</ol>
When fully dissolved, mix solution (a) and (b) 
*Keep away from light*
3. Bisulfite Mutagenesis
<ol>
	<li>Fully remove the 500 &mu;l SDS lysis buffer and add 300 &mu;l mineral oil (~20 hours). Any lysis buffer remaining will dilute the agarose when it is heated and the bead will be more susceptible to dissolving in the subsequent steps. Proceed with bisulfite mutagenesis immediately, or store at -20 &deg;C for up to 5 days.</li>
	<li>If applicable, remove oocytes from the freezer and let thaw (only until agarose bead is relatively translucent). Incubate for 2.5 minutes on a 90 &deg;C heat block, following which Incubate on ice for 10 minutes. 
	Note: Do not mix or stir, extend longer than 2.5 minutes, or fluctuate temperature.</li>
	<li>To perform denaturation, remove the mineral oil and add 1 ml 0.1M NaOH to each tube, flick and invert 5-6 times.</li>
	<li>Incubate for 15 minutes in a 37 &deg;C waterbath, inverting every 3-4 minutes. The bead should float in the NaOH.</li>
	<li>To perform bisulfite treatment, spin the tube gently, then remove the NaOH and add 300 &mu;l mineral oil and 500 &mu;l bisulfite solution. Incubate the tube for 3.5 hours in a 50 &deg;C waterbath. *Keep away from light* 
	Note: Length of incubation may need to be empirically determined for gene of interest.</li>
	<li>To perform desulfonation, incubate on ice for 3 minutes, then remove the mineral oil and the bisulfite solution, spin gently, and add 1 ml 0.3 M NaOH. Flick and invert 5-6 times.</li>
	<li>Incubate for 15 minutes in a 37 &deg;C waterbath, inverting every 3-4 minutes. The bead should float in the NaOH.</li>
	<li>Wash the samples, by first spinning gently, then remove the NaOH and add 1 ml 1x TE pH 7.5. Shake for 5-10 minutes at room temperature (on a shaker). Spin gently again, then remove the 1x TE. Repeat this washing process twice.</li>
	<li>Add 1 ml autoclaved ddH2O. Shake for 5-10 minutes at room temperature (on a shaker). Spin gently, then remove the H2O. Repeat ddH2O wash twice.</li>
	<li>Check the pH of the supernatant; it should be pH 5.0. If still too basic, wash again with H2O. Remove all supernatant, leaving only the agarose bead.</li>
</ol>
4. 1st and 2nd Round PCR amplification
<ol>
	<li>Prepare 1st round PCR mix **while washing**</li>
</ol>
<table border="0">
	<tbody>
		<tr>
			<td>10 &mu;M Primer Forward Outer</td>
			<td>0.5 &mu;l</td>
		</tr>
		<tr>
			<td>10 &mu;M Primer Reverse Outer</td>
			<td>0.5 &mu;l</td>
		</tr>
		<tr>
			<td>240 ng/ml tRNA</td>
			<td>1 &mu;l</td>
		</tr>
		<tr>
			<td>H2O</td>
			<td>13 &mu;l</td>
		</tr>
	</tbody>
</table>
Add to Illustra Ready-to-Go Hot Start PCR beads 
Carefully slide the solid agarose bead into the PCR tube (~10 &mu;l) 
Heat to 70 &deg;C and mix 
Add 25 &mu;l mineral oil 
Total: 50 &mu;l
<ol start="2">
	<li>Amplify 
	Note: An example of cycling conditions for mouse Snrpn is denaturation for 2 minutes at 94 &deg;C, followed by 40 cycles of 30 seconds at 94 &deg;C, 1 minute at 50 &deg;C, and 1 minute at 68 &deg;C; and a final 10 minute elongation step at 68 &deg;C. Annealing temperature for 1st round PCR for mouse H19 and Peg3 is 50 &deg;C.</li>
	<li>Prepare 2nd round PCR mix</li>
</ol>
<table border="0">
	<tbody>
		<tr>
			<td>10 &mu;M Primer Forward Inner</td>
			<td>0.5 &mu;l</td>
		</tr>
		<tr>
			<td>10 &mu;M Primer Reverse Inner</td>
			<td>0.5 &mu;l</td>
		</tr>
		<tr>
			<td>H2O</td>
			<td>19 &mu;l</td>
		</tr>
	</tbody>
</table>
Add to Illustra Ready-to-Go Hot Start PCR beads 
Add 5 &mu;l 1st Round product as a template. Heat the 1st round product to 70 &deg;C for 1 minute to soften the agarose. Be sure to pipette below the layer of mineral oil. 
Add 25 &mu;l mineral oil 
Total: 50 &mu;l
Note: Nested primer sequences for Snrpn, H19, and Peg3 can be found in Market-Velker et al10,12.
<ol start="4">
	<li>Amplify 
	Note: Cycling conditions for mouse Snrpn is denaturation for 2 minutes at 94 &deg;C, followed by 40 cycles of 30 seconds at 94 &deg;C, 1 minute at 50 &deg;C, and 1 minute at 68 &deg;C; and a final 10 minute elongation step at 68 &deg;C10. Mouse H19 and Peg3 require a 50 &deg;C annealing temperature for 2nd round PCR.</li>
	<li>As a diagnostic test, second round samples can be cut with a restriction enzyme that is methylation- or strain-specific.</li>
</ol>
<table border="0">
	<tbody>
		<tr>
			<td>2nd Round product</td>
			<td>4 &mu;l</td>
		</tr>
		<tr>
			<td>Restriction Enzyme</td>
			<td>1 &mu;l</td>
		</tr>
		<tr>
			<td>Buffer</td>
			<td>1 &mu;l</td>
		</tr>
		<tr>
			<td>H2O</td>
			<td>4 &mu;l</td>
		</tr>
	</tbody>
</table>
<ol start="6">
	<li>Electrophorese the digestion products on an 8% acrylamide gel. Any heterogeneous bands represent more than one sequence.</li>
</ol>
5. TA Cloning and Colony PCR
<ol>
	<li>To clone 2nd round product, first heat to 70 &deg;C for 1 minute to soften the agarose, then ligate into vector using the Promega pGEM-T Vector System (Fisher Scientific Cat#A1360).</li>
</ol>
<table border="0">
	<tbody>
		<tr>
			<td>2nd Round PCR</td>
			<td>1 &mu;l</td>
		</tr>
		<tr>
			<td>pGEMT-EASY vector</td>
			<td>1 &mu;l</td>
		</tr>
		<tr>
			<td>Ligase</td>
			<td>1 &mu;l</td>
		</tr>
		<tr>
			<td>H2O</td>
			<td>2 &mu;l</td>
		</tr>
		<tr>
			<td>2x Ligation Buffer</td>
			<td>5 &mu;l</td>
		</tr>
	</tbody>
</table>
Incubate overnight @ 4 &deg;C in PCR machine.
<ol start="2">
	<li>Thaw competent E.coli cells on ice for 15 minutes (Zymo Research Corp Cat#T3009). Add 3 &mu;l ligation reaction to 8 &mu;l E.coli and incubate ligation on ice for 15 minutes.</li>
	<li>Heat shock for 40 seconds in a 42 &deg;C waterbath, and incubate on ice for 2 minutes. Add 60 &mu;l S.O.C. medium and incubate at 37 &deg;C for 1 hour (in shaker).</li>
	<li>Place all of the reaction mix on an LB/Agar/IPTG/Xgal/Amp plate and incubate plate at 37 &deg;C overnight.</li>
	<li>Prepare colony PCR mix</li>
</ol>
<table border="0">
	<tbody>
		<tr>
			<td>20 &mu;M M13 Forward Primer</td>
			<td>0.7 &mu;l</td>
		</tr>
		<tr>
			<td>20 &mu;M M13 Reverse Primer</td>
			<td>0.7 &mu;l</td>
		</tr>
		<tr>
			<td>5X Green Go Taq Buffer</td>
			<td>7.0 &mu;l</td>
		</tr>
		<tr>
			<td>10 mM dNTP</td>
			<td>0.7 &mu;l</td>
		</tr>
		<tr>
			<td>Taq DNA polymerase</td>
			<td>0.28 &mu;l</td>
		</tr>
		<tr>
			<td>H2O</td>
			<td>25.62 &mu;l</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>35 &mu;l Total</td>
		</tr>
	</tbody>
</table>
Add 35 &mu;l Colony PCR master mix into a PCR tube. Pick a white bacterial colony from the plate with a pipette tip, and swirl it into the PCR reaction.
<ol start="6">
	<li>Amplify with denaturation for 10 minutes at 94 &deg;C, followed by 30 cycles of 45 seconds at 94 &deg;C, 30 seconds at 57 &deg;C, and 1 minute at 72 &deg;C; and a final 10 minute elongation step at 72 &deg;C. Electrophorese 4 &mu;l on a 1.5% agarose gel. Send ~30 &mu;l of the PCR product for sequencing. 
	Note: For oocytes, 5 colony PCR products are sequenced.</li>
	<li>Once sequencing results are obtained, methylation patterns can be read. Any original CG that remained as a CG was methylated, and any original CG that is now a TG was unmethylated.</li>
</ol>
6. Representative Results
In our work, we assay imprinted methylation in individual oocytes and embryos (Figure 1). Following nested PCR amplification using bisulfite converted primers, it is possible to confirm a successful conversion by visualizing a correct fragment size on an agarose gel (Figure 2). An individual oocyte represents one parental allele, and in theory, has one imprinted methylation pattern. As such, second round PCR products can be tested for unintentional contamination. A restriction enzyme sensitive to DNA methylation (such as HinfI or DpnII) can be used to digest the second round PCR product to assess whether it contains a methylated or unmethylated allele (Figure 3). A methylated C within the enzyme recognition sequence is cleaved while an unmethylated C that is converted to T is no longer recognized by the enzyme and is uncut. Any MII oocyte sample containing both methylated and unmethylated alleles should be discarded, as it is indicative of cumulus cell contamination (Figure 3). Following ligation and transformation, successful colony PCR amplification can be visualized on an agarose gel to ensure samples with the correct product size are sent for sequencing (Figure 4). Finally, the sequence of five individual clones from an MII oocyte should produce five identical methylation patterns and identical nonCpG conversion rates (Figure 5a). Any samples that contain more than one pattern should be discarded (Figure 5b). Since ovulated MII oocytes have two chromosome copies or an attached polar body, there is a possibility for obtaining two similar sequence patterns (Figure 5c). We recommend discarding data from oocytes that have highly dissimilar methylation patterns since cumulus cell contamination cannot be ruled out.
 
<img alt="Figure 1" src="/files/ftp_upload/4046/4046fig1.jpg" /> 
Figure 1. Schematic of the Single Oocyte Bisulfite Mutagenesis assay.
<img alt="Figure 2" src="/files/ftp_upload/4046/4046fig2.jpg" /> 
Figure 2. Representative results from 2nd round amplification for Snrpn from a single MII oocyte on a 1.5% agarose gel. Lanes 1-4 are four single MII oocytes and lane 5 is a negative control (no oocyte). Expected amplicon size for Snrpn is 420 bp. L, ladder.
<img alt="Figure 3" src="/files/ftp_upload/4046/4046fig3.jpg" /> 
Figure 3. Representative results from 2nd round methylation-specific restriction digestion for Snrpn from a single MII oocyte on an 8% acrylamide gel. HinfI diagnostic restriction digestion shows unmethylated DNA which harbors a T that abolishes the restriction site (420bp, lane 1) or methylated DNA which contains a C within recognition site (cut, 262, 103, and 54 bp, lane 2). Digestion showing both methylated and unmethylated restriction enzyme sites (cut &amp; uncut bands, lane 3) are indicative of cumulus cell contamination. L, ladder.
<img alt="Figure 4" src="/files/ftp_upload/4046/4046fig4.jpg" /> 
Figure 4. Representative results for colony PCR amplification for Snrpn from a single MII oocyte on a 1.5% agarose gel. Expected amplicon size following ligation of Snrpn into the pGEM-T Easy vector and using M13 forward and reverse primers is 656 bp. Lane 1-8, amplicons from clones 1-8. Clone 5 has an incorrect amplicon size and should not be sent for sequencing.
<img alt="Figure 5" src="/files/ftp_upload/4046/4046fig5.jpg" /> 
Figure 5. Representative sequencing results for Snrpn from a single MII oocyte. Snrpn is methylated in oocytes. Black circles indicate methylated CpGs. White circles indicate unmethylated CpGs. CpG number and placement is representative for a B6 strain female mouse. a) Expected sequencing results for Snrpn from a single MII oocyte. Only a single strand of DNA should amplify in all five clones. Oocytes with a single methylation pattern and identical non-CpG conversion pattern should be included in analyses (percent conversion of non-CpGs indicated to the right was calculated as the number of non-CpG cytosines converted to thymine as a percentage of total non-CpG cytosines). b) Sequencing results for Snrpn from a single MII oocyte with cumulus cell contamination. Note the dissimilarity between methylation states and conversion patterns indicating multiple strand amplification. c) Sequencing results for Snrpn from a single MII oocyte with both chromosome copies or polar body inclusion.

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	<li>Jaenisch, R., Bird, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epigenetic+regulation+of+gene+expression:+how+the+genome+integrates+intrinsic+and+environmental+signals.">Epigenetic regulation of gene expression: how the genome integrates intrinsic and environmental signals.</a> Nat. 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The authors have nothing to disclose.

This single oocyte assay contains many steps with a number that are critical and require special care. The first is oocyte washing. It is particularly important to wash each oocyte multiple times in fresh medium drops following hyaluronidase treatment to remove as many cumulus cells as possible. Moreover, when transferring oocytes to acidic tyrode's solution for zona pellucida removal make sure surrounding medium is clear of cumulus cells. The oocyte is very sticky following zona removal, and any surrounding cumulus cell...

Table of specific reagents and equipment.
<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td colspan="4">Oocyte Collection</td>
		</tr>
		<tr>
			<td>Hyaluronidase</td>
			<td>Sigma</td>
			<td>H4272</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Acidic Tyrode</td>
			<td>Sigma</td>
			<td>T1788</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Proteinase K</td>
			<td>Sigma</td>
			<td>P5568</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>10% IGEPAL</td>
			<td>Bioshop</td>
			<td>NON999.500</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td colspan="4">Lysis Solution</td>
		</tr>
		<tr>
			<td>Tris pH 7.5</td>
			<td>Bioshop</td>
			<td>TRS001.5</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>LiCl</td>
			<td>Sigma</td>
			<td>L9650</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>EDTA pH 8.0</td>
			<td>Sigma</td>
			<td>E5134</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>LiDS</td>
			<td>Bioshop</td>
			<td>LDS701.10</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>DTT</td>
			<td>Invitrogen</td>
			<td>P2325</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td colspan="4">SDS Lysis Buffer</td>
		</tr>
		<tr>
			<td>TE pH 7.5</td>
			<td>
			Bioshop(Tris)
			Sigma (EDTA)</td>
			<td>
			TRS001.5
			E5134</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>10% SDS</td>
			<td>Bioshop</td>
			<td>SDS001.500</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td colspan="4">Bisulfite Conversion</td>
		</tr>
		<tr>
			<td>Sodium Hydroxide</td>
			<td>Sigma</td>
			<td>S8045</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Sodium Hydrogensulfite (Sodium Bisulfite)</td>
			<td>Sigma</td>
			<td>243973</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Hydroquinone</td>
			<td>Sigma</td>
			<td>H9003</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Low Melting Point (LMP) Agarose</td>
			<td>Sigma</td>
			<td>A9414</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Mineral Oil</td>
			<td>Sigma</td>
			<td>M8410</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>M2 Medium</td>
			<td>Sigma</td>
			<td>M7167</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>GIBCO Distilled water</td>
			<td>Invitrogen</td>
			<td>15230-196</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Autoclaved double distilled (dd) water</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td colspan="4">PCR</td>
		</tr>
		<tr>
			<td>Illustra Hot Start Mix RTG</td>
			<td>GE Healthcare</td>
			<td>28-9006-54</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>240 ng/ml yeast tRNA</td>
			<td>Invitrogen</td>
			<td>15401-011</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>5x Green GoTaq Reaction Buffer</td>
			<td>Promega</td>
			<td>M7911</td>
			<td>&nbsp;</td>
		</tr>
<tr>
			<td>Inner and outer nested primers</td>
			<td>Sigma</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td colspan="4">Ligation</td>
		</tr>
		<tr>
			<td>Promega pGEM-T Easy Vector</td>
			<td>Fisher Scientific</td>
			<td>A1360</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td colspan="4">TA Cloning</td>
		</tr>
		<tr>
			<td>Competent E.coli cells</td>
			<td>Zymo Research Corp.</td>
			<td>T3009</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td colspan="4">Equipment</td>
		</tr>
		<tr>
			<td>Dissecting Microscope</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>70&deg;C and 90&deg;C Heat Blocks</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>37&deg;C and 50&deg;C Waterbaths (42&deg;C for transformations)</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Rocker</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>PCR machine</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

single oocyte bisulfite mutagenesis

Epigenetics encompasses all heritable and reversible modifications to chromatin that alter gene accessibility, and thus are the primary mechanisms for regulating gene transcription1. DNA methylation is an epigenetic modification that acts predominantly as a repressive mark. Through the covalent addition of a methyl group onto cytosines in CpG dinucleotides, it can recruit additional repressive proteins and histone modifications to initiate processes involved in condensing chromatin and silencing genes2. DNA methylation is essential for normal development as it plays a critical role in developmental programming, cell differentiation, repression of retroviral elements, X-chromosome inactivation and genomic imprinting.
One of the most powerful methods for DNA methylation analysis is bisulfite mutagenesis. Sodium bisulfite is a DNA mutagen that deaminates cytosines into uracils. Following PCR amplification and sequencing, these conversion events are detected as thymines. Methylated cytosines are protected from deamination and thus remain as cytosines, enabling identification of DNA methylation at the individual nucleotide level3. Development of the bisulfite mutagenesis assay has advanced from those originally reported4-6 towards ones that are more sensitive and reproducible7. One key advancement was embedding smaller amounts of DNA in an agarose bead, thereby protecting DNA from the harsh bisulfite treatment8. This enabled methylation analysis to be performed on pools of oocytes and blastocyst-stage embryos9. The most sophisticated bisulfite mutagenesis protocol to date is for individual blastocyst-stage embryos10. However, since blastocysts have on average 64 cells (containing 120-720 pg of genomic DNA), this method is not efficacious for methylation studies on individual oocytes or cleavage-stage embryos.
Taking clues from agarose embedding of minute DNA amounts including oocytes11, here we present a method whereby oocytes are directly embedded in an agarose and lysis solution bead immediately following retrieval and removal of the zona pellucida from the oocyte. This enables us to bypass the two main challenges of single oocyte bisulfite mutagenesis: protecting a minute amount of DNA from degradation, and subsequent loss during the numerous protocol steps. Importantly, as data are obtained from single oocytes, the issue of PCR bias within pools is eliminated. Furthermore, inadvertent cumulus cell contamination is detectable by this method since any sample with more than one methylation pattern may be excluded from analysis12. This protocol provides an improved method for successful and reproducible analyses of DNA methylation at the single-cell level and is ideally suited for individual oocytes as well as cleavage-stage embryos.

Watch this Scientific Journal Video about Single Oocyte Bisulfite Mutagenesis at JoVE.com

Single Oocyte Bisulfite Mutagenesis

Bisulfite mutagenesis is the gold standard for analyzing DNA methylation. Our modified protocol allows for DNA methylation analysis at the single-cell level and was specifically designed for individual oocytes. It can also be used for cleavage-stage embryos.

Epigenetics encompasses all heritable and reversible modifications to chromatin that alter gene accessibility, and thus are the primary mechanisms for ...

Research

JoVE Journal

Biology

14.0K Views.  Schulich School of Medicine and Dentistry, University of Western Ontario. Bisulfite mutagenesis is the gold standard for analyzing DNA methylation. Our modified protocol allows for DNA methylation analysis at the single-cell level and was specifically designed for individual oocytes. It can also be used for cleavage-stage embryos.

Video: Single Oocyte Bisulfite Mutagenesis

Homemade Site Directed Mutagenesis of Whole Plasmids

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

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

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

Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying potassium (referred to as GIRK) channels, which are important for regulating the excitability of neurons. There are four different mammalian GIRK channel subunits (GIRK1-GIRK4) - we focus on GIRK2 because it forms a homotetramer. Stimulation of different types of G protein-coupled receptors (GPCRs), such as the muscarinic receptor (M2R), leads to activation of GIRK channels. Alcohol also directly activates GIRK channels. We will show how to mutate one amino acid by specifically changing one or more nucleotides in the cDNA for the GIRK channel. This mutated cDNA sequence will be amplified in bacteria, purified, and the presence of the point mutation will be confirmed by DNA sequencing. The cDNAs for the mutated and wild-type GIRK channels will be transfected into human embryonic kidney HEK293T cells cultured in vitro. Lastly, whole-cell patch-clamp electrophysiology will be used to study the macroscopic potassium currents through the ectopically expressed wild-type or mutated GIRK channels. In this experiment, we will examine the effect of a L257W mutation in GIRK2 channels on M2R-dependent and alcohol-dependent activation.

We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying ...

An Introduction to Worm Lab: from Culturing Worms to Mutagenesis

This protocol describes procedures to maintain nematodes in the laboratory and how to mutagenize them using two alternative methods: ethyl methane sulfonate (EMS) and 4, 5', 8-trimethylpsoralen combined with ultraviolet light (TMP/UV). Nematodes are powerful biological systems for genetics studies because of their simple body plan and mating system, which is composed of self-fertilizing hermaphrodites and males that can generate hundreds of progeny per animal. Nematodes are maintained in agar plates containing a lawn of bacteria and can be easily transferred from one plate to another using a pick. EMS is an alkylating agent commonly used to induce point mutations and small deletions, while TMP/UV mainly induces deletions. Depending on the species of nematode being used, concentrations of EMS and TMP will have to be optimized. To isolate recessive mutations of the nematode Pristionchus pacificus, animals of the F2 generation were visually screened for phenotypes. To illustrate these methods, we mutagenized worms and looked for Uncoordinated (Unc), Dumpy (Dpy) and Transformer (Tra) mutants.

Screening for mutants with phenotypic defects is a straightforward method for identifying genes that function in a given biological process. In this article we describe how to culture free living worms (e.g., Pristionchus pacificus) in the laboratory and show two different mutagenesis methods, EMS and TMP/UV.

This protocol describes procedures to maintain nematodes in the laboratory and how to mutagenize them using two alternative methods: ethyl methane ...

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular signatures, or to evolve proteins in the laboratory. Here, we present a protocol that allows the generation of large (>1011) mutant libraries for a given target sequence. This method is based on replication of a ColE1 plasmid encoding the desired sequence by a low-fidelity variant of DNA polymerase I (LF-Pol I). The target plasmid is transformed into a mutator strain of E. coli and plated on solid media, yielding between 0.2 and 1 mutations/kb, depending on the location of the target gene. Higher mutation frequencies are achieved by iterating this process of mutagenesis. Compared to alternative methods of mutagenesis, our protocol stands out for its simplicity, as no cloning or PCR are involved. Thus, our method is ideal for mutational labeling of plasmids or other Pol I templates or to explore large sections of sequence space for the evolution of activities not present in the original target. The tight spatial control that PCR or randomized oligonucleotide-based methods offer can also be achieved through subsequent cloning of specific sections of the library. Here we provide protocols showing how to create a random mutant library and how to establish drug-based selections in E. coli to identify mutants exhibiting new biochemical activities.

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

Here we demonstrate a simple protocol to create a random mutant library for a given target sequence. We show how this method, which is performed in vivo in Escherichia coli, can be coupled with functional selections to evolve new enzymatic activities.

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular ...

Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

Mistakes in chromosome segregation lead to aneuploid cells. In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to infertility, miscarriages or developmental disorders like Down syndrome. Haploid gametes form through species-specific developmental programs that are coupled to meiosis. The first meiotic division (MI) is unique to meiosis because sister chromatids remain attached while homologous chromosomes are segregated. For reasons not fully understood, this reductional division is prone to errors and is more commonly the source of aneuploidy than errors in meiosis II (MII) or than errors in male meiosis 1,2. 
In mammals, oocytes arrest at prophase of MI with a large, intact germinal vesicle (GV; nucleus) and only resume meiosis when they receive ovulatory cues. Once meiosis resumes, oocytes complete MI and undergo an asymmetric cell division, arresting again at metaphase of MII. Eggs will not complete MII until they are fertilized by sperm. Oocytes also can undergo meiotic maturation using established in vitro culture conditions 3. Because generation of transgenic and gene-targeted mouse mutants is costly and can take long periods of time, manipulation of female gametes in vitro is a more economical and time-saving strategy. 
Here, we describe methods to isolate prophase-arrested oocytes from mice and for microinjection. Any material of choice may be introduced into the oocyte, but because meiotically-competent oocytes are transcriptionally silent 4,5 cRNA, and not DNA, must be injected for ectopic expression studies. To assess ploidy, we describe our conditions for in vitro maturation of oocytes to MII eggs. Historically, chromosome-spreading techniques are used for counting chromosome number 6. This method is technically challenging and is limited to only identifying hyperploidies. Here, we describe a method to determine hypo-and hyperploidies using intact eggs 7-8. This method uses monastrol, a kinesin-5 inhibitor, that collapses the bipolar spindle into a monopolar spindle 9 thus separating chromosomes such that individual kinetochores can readily be detected and counted by using an anti-CREST autoimmune serum. Because this method is performed in intact eggs, chromosomes are not lost due to operator error.

Oocytes are prone to aneuploidy due to errors in chromosome segregation during meiotic maturation. Aneuploid eggs can cause infertility, miscarriages or developmental disorders like Down syndrome. Here, we describe methods to introduce materials of choice into oocytes and methods to study meiotic maturation and assess ploidy.

Mistakes in chromosome segregation lead to aneuploid cells.  In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to ...

Yeast Luminometric and Xenopus Oocyte Electrophysiological Examinations of the Molecular Mechanosensitivity of TRPV4

TRPV4 (Transient Receptor Potentials, vanilloid family, type 4) is widely expressed in vertebrate tissues and is activated by several stimuli, including by mechanical forces. Certain TRPV4 mutations cause complex hereditary bone or neuronal pathologies in human. Wild-type or mutant TRPV4 transgenes are commonly expressed in cultured mammalian cells and examined by Fura-2 fluorometry and by electrodes. In terms of the mechanism of mechanosensitivity and the molecular bases of the diseases, the current literature is confusing and controversial. To complement existing methods, we describe two additional methods to examine the molecular properties of TRPV4. (1) Rat TRPV4 and an aequorin transgene are transformed into budding yeast. A hypo-osmtic shock of the transformant population yields a luminometric signal due to the combination of aequorin with Ca2+, released through the TRPV4 channel. Here TRPV4 is isolated from its usual mammalian partner proteins and reveals its own mechanosensitivity. (2) cRNA of TRPV4 is injected into Xenopus oocytes. After a suitable period of incubation, the macroscopic TRPV4 current is examined with a two-electrode voltage clamp. The current rise upon removal of inert osmoticum from the oocyte bath is indicative of mechanosensitivity. The microAmpere (10-6 to 10-4 A) currents from oocytes are much larger than the subnano- to nanoAmpere (10-10 to 10-9 A) currents from cultured cells, yielding clearer quantifications and more confident assessments. Microscopic currents reflecting the activities of individual channel proteins can also be directly registered under a patch clamp, in on-cell or excised mode. The same oocyte provides multiple patch samples, allowing better data replication. Suctions applied to the patches can activate TRPV4 to directly assess mechanosensitivity. These methods should also be useful in the study of other types of TRP channels.

Yeast Luminometric and Xenopus Oocyte Electrophysiological Examinations of the Molecular Mechanosensitivity of TRPV4

To complement commonly used methods to study TRPV4&#8217;s, two methods are described: Its mechanosensitivity can be studied by Ca2+-aequorin luminometry in transgenic yeast upon hypo-osmotic challenge. It can also be examined in TRPV4-RNA injected Xenopus oocytes by whole-cell two-electrode voltage clamp or patch clamp in on-cell or excised mode.

TRPV4 (Transient Receptor Potentials, vanilloid family, type 4) is widely expressed in vertebrate tissues and is activated by several stimuli, including ...

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion channels in oocytes. Recordings from the cut-open setup are particularly useful for resolving low magnitude gating currents, rapid ionic current activation, and deactivation. The main benefits over the two-electrode voltage clamp (TEVC) technique include increased clamp speed, improved signal-to-noise ratio, and the ability to modulate the intracellular and extracellular milieu.
Here, we employ the human cardiac sodium channel (hNaV1.5), expressed in Xenopus oocytes, to demonstrate the cut-open setup and protocol as well as modifications that are required to add voltage clamp fluorometry capability.
The properties of fast activating ion channels, such as hNaV1.5, cannot be fully resolved near room temperature using TEVC, in which the entirety of the oocyte membrane is clamped, making voltage control difficult. However, in the cut-open technique, isolation of only a small portion of the cell membrane allows for the rapid clamping required to accurately record fast kinetics while preventing channel run-down associated with patch clamp techniques.
In conjunction with the COVG technique, ion channel kinetics and electrophysiological properties can be further assayed by using voltage clamp fluorometry, where protein motion is tracked via cysteine conjugation of extracellularly applied fluorophores, insertion of genetically encoded fluorescent proteins, or the incorporation of unnatural amino acids into the region of interest1. This additional data yields kinetic information about voltage-dependent conformational rearrangements of the protein via changes in the microenvironment surrounding the fluorescent molecule.

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open Vaseline gap approach is used to obtain low noise recordings of ionic and gating currents from voltage-dependent ion channels expressed in Xenopus oocytes with high resolution of fast channel kinetics. With minor modification, voltage clamp fluorometry can be coupled to the cut-open oocyte protocol.

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion ...

Generation of Enterobacter sp. YSU Auxotrophs Using Transposon Mutagenesis

Prototrophic bacteria grow on M-9 minimal salts medium supplemented with glucose (M-9 medium), which is used as a carbon and energy source. Auxotrophs can be generated using a transposome. The commercially available, Tn5-derived transposome used in this protocol consists of a linear segment of DNA containing an R6K&#947; replication origin, a gene for kanamycin resistance and two mosaic sequence ends, which serve as transposase binding sites. The transposome, provided as a DNA/transposase protein complex, is introduced by electroporation into the prototrophic strain, Enterobacter sp. YSU, and randomly incorporates itself into this host&#8217;s genome. Transformants are replica plated onto Luria-Bertani agar plates containing kanamycin, (LB-kan) and onto M-9 medium agar plates containing kanamycin (M-9-kan). The transformants that grow on LB-kan plates but not on M-9-kan plates are considered to be auxotrophs. Purified genomic DNA from an auxotroph is partially digested, ligated and transformed into a pir+ Escherichia coli (E. coli) strain. The R6K&#947; replication origin allows the plasmid to replicate in pir+ E. coli strains, and the kanamycin resistance marker allows for plasmid selection. Each transformant possesses a new plasmid containing the transposon flanked by the interrupted chromosomal region. Sanger sequencing and the Basic Local Alignment Search Tool (BLAST) suggest a putative identity of the interrupted gene. There are three advantages to using this transposome mutagenesis strategy. First, it does not rely on the expression of a transposase gene by the host. Second, the transposome is introduced into the target host by electroporation, rather than by conjugation or by transduction and therefore is more efficient. Third, the R6K&#947; replication origin makes it easy to identify the mutated gene which is partially recovered in a recombinant plasmid. This technique can be used to investigate the genes involved in other characteristics of Enterobacter sp. YSU or of a wider variety of bacterial strains.

Generation of Enterobacter sp. YSU Auxotrophs Using Transposon Mutagenesis

Enterobacter sp. YSU grows in glucose minimal salts medium. Auxotrophs are generated by transforming it with a transposome which randomly inserts itself into the host genome. Mutants are found by replica plating from complex medium to minimal medium. Interrupted genes are identified by gene rescue and sequencing.

Prototrophic bacteria grow on M-9 minimal salts medium supplemented with glucose (M-9 medium), which is used as a carbon and energy source. Auxotrophs can ...

Enhanced Reduced Representation Bisulfite Sequencing for Assessment of DNA Methylation at Base Pair Resolution

DNA methylation pattern mapping is heavily studied in normal and diseased tissues. A variety of methods have been established to interrogate the cytosine methylation patterns in cells. Reduced representation of whole genome bisulfite sequencing was developed to detect quantitative base pair resolution cytosine methylation patterns at GC-rich genomic loci. This is accomplished by combining the use of a restriction enzyme followed by bisulfite conversion. Enhanced Reduced Representation Bisulfite Sequencing (ERRBS) increases the biologically relevant genomic loci covered and has been used to profile cytosine methylation in DNA from human, mouse and other organisms. ERRBS initiates with restriction enzyme digestion of DNA to generate low molecular weight fragments for use in library preparation. These fragments are subjected to standard library construction for next generation sequencing. Bisulfite conversion of unmethylated cytosines prior to the final amplification step allows for quantitative base resolution of cytosine methylation levels in covered genomic loci. The protocol can be completed within four days. Despite low complexity in the first three bases sequenced, ERRBS libraries yield high quality data when using a designated sequencing control lane. Mapping and bioinformatics analysis is then performed and yields data that can be easily integrated with a variety of genome-wide platforms. ERRBS can utilize small input material quantities making it feasible to process human clinical samples and applicable in a range of research applications. The video produced demonstrates critical steps of the ERRBS protocol.

Enhanced Reduced Representation Bisulfite Sequencing is a method for the preparation of sequencing libraries for DNA methylation analysis based on restriction enzyme digestion combined with cytosine bisulfite conversion. This protocol requires 50 ng of starting material and yields base pair resolution data at GC-rich genomic regions.

DNA methylation pattern mapping is heavily studied in normal and diseased tissues. A variety of methods have been established to interrogate the cytosine ...

A Protocol for Functional Assessment of Whole-Protein Saturation Mutagenesis Libraries Utilizing High-Throughput Sequencing

Site-directed mutagenesis has long been used as a method to interrogate protein structure, function and evolution. Recent advances in massively-parallel sequencing technology have opened up the possibility of assessing the functional or fitness effects of large numbers of mutations simultaneously. Here, we present a protocol for experimentally determining the effects of all possible single amino acid mutations in a protein of interest utilizing high-throughput sequencing technology, using the 263 amino acid antibiotic resistance enzyme TEM-1 &#946;-lactamase as an example. In this approach, a whole-protein saturation mutagenesis library is constructed by site-directed mutagenic PCR, randomizing each position individually to all possible amino acids. The library is then transformed into bacteria, and selected for the ability to confer resistance to &#946;-lactam antibiotics. The fitness effect of each mutation is then determined by deep sequencing of the library before and after selection. Importantly, this protocol introduces methods which maximize sequencing read depth and permit the simultaneous selection of the entire mutation library, by mixing adjacent positions into groups of length accommodated by high-throughput sequencing read length and utilizing orthogonal primers to barcode each group. Representative results using this protocol are provided by assessing the fitness effects of all single amino acid mutations in TEM-1 at a clinically relevant dosage of ampicillin. The method should be easily extendable to other proteins for which a high-throughput selection assay is in place.

We present a protocol for the functional assessment of comprehensive single-site saturation mutagenesis libraries of proteins utilizing high-throughput sequencing. Importantly, this approach uses orthogonal primer pairs to multiplex library construction and sequencing. Representative results using TEM-1 &#946;-lactamase selected at a clinically relevant dosage of ampicillin are provided.

Site-directed mutagenesis has long been used as a method to interrogate protein structure, function and evolution. Recent advances in massively-parallel ...