We thank members in the Zhang lab for providing stimulating discussions and members in the Zhan lab for providing technical assistance. This research was supported by grants 31500860 (to C.Z.) of the NSFC, 2012CB837700 (to E.E.Z. and C.Z.) of the 973 Program from the M.O.S.T. of China, and by funding from the Beijing Municipal Government. E.E.Z. was supported by the Chinese &#34;Recruitment Program of Global Youth Experts&#34;.

All procedures in this protocol were conducted with the approval of the Institutional Animal Care and Use Committee (IACUC) of the National Institute of Biological Sciences, Beijing, in accordance with the governmental regulations of China.
1. Construction of the Cry1 Fluorescence Reporter
Note: Previous circadian studies using the bioluminescence system2,12,13 fuse a circadian promoter with a destabilized luciferase (dLuc) to induce rhythmic dLuc expression. To develop a fluorescence reporter, destabilized luciferase was replaced with destabilized fluorescence protein (dVenus) to induce rhythmic dVenus expression. The generation of the P(Cry1)-dVenus reporter is described below, the construct of which is currently being submitted to Addgene and will be available soon for academic use.
<ol>
	<li>Amplify the functional mouse Cry1 promoter (+328 to -1208 bp region of the Cry1 gene, transcription start site designated as &#8216;&#8216;-1&#8217;&#8217;). Use the PCR enzyme (KOD Plus Neo), primers F1 and R1 (see Table 2), mouse genome as a template and the manufacturer&#8217;s PCR protocol to amplify the Cry1 promoter DNA fragment.</li>
	<li>Amplify intron 336 of the Cry1 gene, which is important for the circadian clock function of Cry114. Use the PCR enzyme, primers F2 and R2, mouse genome as a template and the manufacturer&#8217;s PCR protocol to amplify the intron 336 DNA fragment.</li>
	<li>Amplify Venus. Use the PCR enzyme, primers F3 and R3, pVENUS-N1 plasmid as a template and the manufacturer&#8217;s PCR protocol to amplify the Venus DNA fragment.</li>
	<li>Amplify the NLS-D2 DNA fragment. Use the PCR enzyme, primers F4 and R4, pcDNA3.3-d2eGFP plasmid as a template and the manufacturer&#8217;s PCR protocol to amplify the NLS-D2 DNA fragment.</li>
	<li>Synthesize DNA fragment 1 as exon 1.</li>
	<li>Synthesize DNA fragment 2 as the linker between intron 336 and the Venus DNA fragment.</li>
	<li>Cut the pAAV-EF1a-double floxed-hChR2(H134R)-mCherry-WPRE-HGHpA vector plasmid with the MluI and EcoRI enzymes. Use the Gel Extraction Kit to purify the larger DNA fragment. Use the manufacturer&#8217;s protocol.</li>
	<li>Using a Gibson assembly mix and its standard protocol, assemble all of the DNA fragments together to produce the pAAV-P(Cry1)-intron336-Venus-NLS-D2-WPRE-HGHpA reporter. Use the manufacturer&#8217;s protocol.</li>
</ol>
2. Production of Adeno-associated Virus15
<ol>
	<li>Prepare the following plasmid DNA stocks: 31.25 &#181;g of pRV1, 31.25 &#181;g of pH21, 125 &#181;g of pFdelta6, and 62.5 &#181;g of pAAV-P(Cry1)-intron336-Venus-NLS- D2-WPRE-HGHpA.</li>
	<li>Prepare five 15 cm dishes of 293T cells and wait until the cells become 90% confluent.</li>
	<li>Transfect the plasmids into the 297T cells following the transfection reagent protocol. For each dish, use 6.25 &#181;g of pRV1, 6.25 &#181;g of pH21, 25 &#181;g of pFdelta6, 12.5 &#181;g of pAAV-P(Cry1)-intron336-Venus-NLS-D2-WPRE-HGHpA, and 120 &#181;L of transfection reagent.</li>
	<li>72 h after transfection, detach the cells from each plate using a pipette gun; harvest the cells into two 50 mL tubes.</li>
	<li>Pellet the cells at 800 x g for 10 min and discard the supernatant. Wash the cells by placing 20 mL of phosphate-buffered saline (PBS) into each tube; pellet the cells at 800 x g for 10 min, and discard the supernatant.</li>
	<li>Resuspend pellets in 150 mM NaCl and 20 mM Tris solution, pH 8.0; use 25 mL of solution/tube. Add 1.25 mL of freshly prepared 10% sodium deoxycholate (in dH2O) to each tube. Add benzonase nuclease to a final concentration of 50 units per mL, and mix thoroughly. Incubate at 37 &#176;C for 1 h.</li>
	<li>Remove cellular debris by centrifuging at 3000 x g for 15 min. Further ensure that all debris has been removed by pushing the mixture through a 0.45 &#956;m syringe filter; this step will help prevent blocking of heparin columns.</li>
	<li>Set up the heparin columns using a peristaltic pump; set the flow rate to 1 mL/min, ensuring that no air bubbles are introduced into the columns.</li>
	<li>Equilibrate the columns with 10 mL of a 150 mM NaCl, 20 mM Tris solution, pH 8.0. Add the solution containing the virus to each column. Wash the columns with 20 mL of a 100 mM NaCl, 20 mM Tris solution, pH 8.0. Then, using a 5 mL syringe, wash the columns with 1 mL of a 200 mM NaCl and 20 mM Tris solution pH 8.0, followed by 1 mL of a 300 mM NaCl and 20 mM Tris solution, pH 8.0.</li>
	<li>Use a 5 mL syringe to elute the virus with 3 mL of a 400 mM NaCl, 20 mM Tris solution, pH 8.0, followed by 4 mL of a 450 mM NaCl, 20 mM Tris solution, pH 8.0, and finally by a 3 mL of a 500 mM NaCl, 20 mM Tris solution, pH 8.0. Collect the extracted materials.</li>
	<li>For recombinant adeno-associated virus (rAAV) concentration, load the extracted materials, 4 mL at a time, into centrifugal filter units with a 100,000 molecular weight cutoff. Centrifuge each 4-mL sample at 2000 x g for 5 min at room temperature, discarding the flow-through.</li>
	<li>When all extracted materials have been run through the centrifugal filter units, add 4 mL of PBS and centrifuge the mixture into approximately 250 &#956;L. Remove rAAVs (viral titer should be about 2 - 8 x 1012 viral particles per mL) from the centrifugal filter units, aliquot and store at -80 &#176;C until use. At this stage, the virus can be stored for several months.</li>
</ol>
3. rAAV Injection and Optic Fiber Insertion into the Adult Mouse SCN
<ol>
	<li>Injection of circadian fluorescence reporter-carrying rAAVs into the SCN
	<ol>
		<li>Load 5 &#181;L of rAAV (the virus titer is about 2 - 8 x 1012 virus particles per mL) into a micro-syringe.</li>
		<li>Anesthetize an adult mouse (2- to 5-month-old, C57BL/6 background) with Nembutal (80 mg/kg) via intraperitoneal injection. Check for sufficient depth of anesthesia by the lack of a toe-pinch response.</li>
		<li>Use scissors to remove fur from the head of the mouse and then mount the mouse into a stereotaxic apparatus. Provide a heat source to anesthetized animals via a circulating hot water blanket or another heating device. 
		NOTE: Alternatively, use depilatory cream or electric hair clippers to remove the fur.</li>
		<li>Clean the surgical site with the appropriate disinfecting agents. For example, this could include sterile water or alcohol and iodine, betadine,&#160;or dilute chlorhexidine in alternating scrubs. Repeat at least 3 times.&#160;Drape the surgical site.</li>
		<li>&#160;Use the scissors to make an incision in the scalp and expose the skull.</li>
		<li>Adjust the stereotaxic apparatus so that bregma and lambda are in the same horizontal plane; make two symmetrical points of bregma in the same horizontal plane.</li>
		<li>Use a sterile cotton swab soaked in 4% H2O2/H2O to corrode the brain periosteum. Then use a clean, sterile cotton swab to clear and dry the surface of the skull.</li>
		<li>Using a micro drill, make a hole about 1 mm in diameter 0.46 mm posterior and 0.25 mm lateral to bregma.</li>
		<li>fiUse a clean cotton sterile swab with physiological saline solution to clear bone fragments from the exposed surface of the brain.</li>
		<li>Insert a micro-syringe into the brain, ensuring that the tip of the micro-syringe is 0.46 mm posterior and 0.25 mm lateral to bregma, and 5.7 mm deep from the surface of the skull.</li>
		<li>Inject 500 nL of rAAV (50 nL min-1) into the SCN, leaving the micro-syringe in place for 10 min after the injection to ensure complete diffusion of rAAV.</li>
		<li>Slowly withdraw the micro-syringe.</li>
	</ol>
	</li>
	<li>Insert the optical fiber into the SCN area.
	<ol>
		<li>Using an optical fiber knife, cut a fiber 17-mm in length, making sure that each side of the optic fiber is smooth.</li>
		<li>Insert the optic fiber into the ceramic ferrule and fix the optic fiber to a ceramic ferrule with methyl ethyl ketone peroxide (AB glue), ensuring that the surfaces of the optical fiber and ceramic ferrule are flush.</li>
		<li>Clean the fiber with a cold sterilant solution or ethylene oxide.</li>
		<li>After rAAV injection (step 3.1.11), insert the ceramic ferrule-containing optic fiber into the SCN.</li>
		<li>Fix the ceramic ferrule and optical fiber onto the skull using dental resin and allow to dry.</li>
		<li>Smear the surface of the dental resin with back nail polish. Repeat the procedures outlined in 3.1.2&#160;to 3.2.5, omitting the actual injection of rAAV for control.</li>
	</ol>
	</li>
	<li>Post-surgical care
	<ol>
		<li>Provide mice with post-operative analgesics, such as buprenorphine 2 mg/kg body weight, subcutaneously, twice a day.</li>
		<li>Place mice in a recovery cage while they come out of anesthesia.</li>
		<li>House mice individually under a 12 h light/dark condition (light intensity ~100 lux at the bottom of the cage) with food/water available ad libitum. Eight days should be allotted before further experimentation to allow for sufficient recovery time and to ensure optimal fluorescence reporter expression.</li>
	</ol>
	</li>
</ol>
4. In vivo Fluorescence Signal Monitoring and Data Collection
<ol>
	<li>Eight days after surgery, connect the fiber on the mouse head with the fluorescence signal monitor (e.g., Gene Observer). Check the fluorescence signal in the SCN of control mice to establish the background signal. Conduct the observation and measurement of this signal using fluorescence signal monitor.</li>
	<li>Evaluate the fluorescence signal of experimental mouse with the same operation of the control mouse. Exclude mice which have received the viral vector but which only display the background signal from analysis, as this likely indicates that the tip of the optic fiber has missed the SCN. After these initial measures, return mice to their home cages for another 3 weeks to allow the fluorescence signal to stabilize.</li>
	<li>After 3 weeks, connect the mice with the fluorescence signal monitor. Measure the fluorescence for 15 s every 10 min, at a frequency of 100 Hz. During these measurements, the mice are freely moving in their cages with food/water available ad libitum. Use a light intensity at the bottom of the cage of ~100 lux, and a the laser power at the tip of the optic fiber between 15 - 20 &#181;W.</li>
	<li>After the completion of all recordings, perfuse mice with 4% paraformaldehyde, and remove and slice the brains to check placement of the optic fiber. Exclude mice from analysis if the optic fiber tip is not correctly implanted in the SCN.</li>
</ol>
5. Data Analysis and Presentation
<ol>
	<li>Measure fluorescence signal every 10 min for 15 s, at a frequency of 100 Hz, to yield 1500 data points. The average of these 1500 points translates to the fluorescence signal at any given time point. Plotting multiple averages over time produces a signal-to-time curve which corresponds to the transcriptional rhythm of Cry1 under a control light condition in the SCN of freely moving mice.</li>
</ol>

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The authors have no conflicts of interest to disclose.

In contrast to ex vivo methods, such as slice culture4,5, RT-PCR16, and in situ hybridization17, which require that animals be killed, the in vivo recording method allows investigators to study circadian gene expression in a living animal. As such, this technology provides the ability to evaluate the effect of different physical perturbations (e.g., sleep deprivation, stress, food intake, etc.) on the neural circadian clock. In contrast to the in vivo bioluminescence7,8,10 method that can only work in constant dark conditions, the fluorescence in vivo recording method can be applied under light conditions-a key advantage of the technology as light signals re-entrainment of the SCN circadian clock18. Although relatively constant, robust circadian rhythms can be detected with this method, it should be mentioned that the reporter technology does not provide quantitative information and can therefore not be used to measure levels of gene transcription and translation.
There are many critical steps of this protocol. The concentrated rAAV titer should be as high as 2 - 8x 1012 viral particles per mL; lower viral titers would give off dim or even undetectable fluorescence signals. When fixing the head of the mouse into the stereotaxic apparatus, ensure that the skull is horizontal to secure accurate positioning for virus injection and optical fiber implantation. Injection of the virus should be done slowly (&#8804;50 nL min-1) to ensure that the virus remains in the SCN. The ceramic ferrule and optical fiber should be securely fixed on the skull and dental resin should not contact the skin. If the ceramic ferrule and optical fiber are not adequately fixed, they will gradually detach from the mouse head.
This technique can be extended to include other gene reporters (e.g., Per2, Bmal1, c-Fos, Pomc, etc.) by changing the promoter and the corresponding intron19. By reversing the flag-inton336-Venus-NLS-D2 cassette with two reversed lox sequences and combining the reporter with a Cre mouse line, this technology can be used to record activity in specific neurons. For example, combining this approach with a Vip-cre mouse line would allow for the recording of Cry1 transcriptional rhythms in vasoactive intestinal polypeptide-expressing neurons in the SCN. This method can also employ GCaMP6 as a Ca2+ indicator to record the Ca2+ circadian rhythm in the brain.

In mammals, the suprachiasmatic nucleus (SCN) governs the whole body&#39;s circadian rhythm to coordinate an individual&#39;s response to exogenous environmental changes (e.g., light, temperature, stress, etc.)1. Core clock components consist of Per1-3, Cry1-2, Clock, and Bmal1, and play a central role in regulating the circadian clock of each cell. Each cell in the SCN contains the transcriptional activator, CLOCK/BMAL1, which acts as a heterodimer to induce the expression of PER and CRY. The PER/CRY complex then inhibits the function of CLOCK/BMAL1 to form a transcription-translation feedback loop that takes about 24 h to complete2,3.
Previous studies on the SCN have mainly employed the ex vivo SCN slice culture method4,5,6 and, while this approach has provided valuable information, its limitations have inhibited our ability to obtain data regarding the influence of other brain nuclei on the SCN, as well as the effect of exogenous stimuli (e.g., light) on cells residing in this critical region. In 2001, Hitoshi Okamura&#39;s group was the first to use the bioluminescence system to in vivo monitor circadian clock gene expression in the SCN in freely moving mice7. Ken-ichi Honma&#39;s group has spent the past few years further developing the bioluminescence in vivo recording system in the SCN8,9,10. Together, these studies have provided researchers with the ability to monitor the circadian clock in constant darkness or after a light pulse. However, because bioluminescence is too dim to allow for continuous monitoring during the light/dark cycle, coupled with the fact that light is the predominant signal required for the SCN-mediated entrainment of circadian clocks11, there is increasing demand for the development of experimental methods which overcome the limitations associated with bioluminescence recording. The current report describes a fluorescence-based system which was constructed to monitor the circadian clock of the SCN in vivo in freely moving mice. This easy-to-use method permits continuous monitoring during the light/dark cycle and allows for the long-term observation of the transcription of circadian clock genes in the SCN in real-time and at high temporal resolution.

<table border="0" cellpadding="0" cellspacing="0" style="width:1408px;" width="1408">
	<tbody>
		<tr height="19">
			<td height="19" style="height:19px;width:627px;">KOD Plus Neo</td>
			<td style="width:403px;">TOYOBO</td>
			<td style="width:133px;">KOD-401</td>
			<td style="width:245px;">Reagent</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">pVENUS-N1</td>
			<td>addgene</td>
			<td>#61854</td>
			<td>Plasmid</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">pcDNA3.3_d2eGFP</td>
			<td>addgene</td>
			<td>#26821</td>
			<td>Plasmid</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:21px;">pAAV-EF1a-double floxed-hChR2(H134R)-mCherry-WPRE-HGHpA</td>
			<td>addgene</td>
			<td>#20297</td>
			<td>Plasmid</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">MluI</td>
			<td>Thermo Scientific</td>
			<td>FD0564</td>
			<td>Reagent</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">EcoRI</td>
			<td>Thermo Scientific</td>
			<td>FD0274</td>
			<td>Reagent</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">Gibson Assembly Mix</td>
			<td>NEB</td>
			<td>E2611s</td>
			<td>Reagent</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">Lipofectamine 2000</td>
			<td>Thermo Scientific</td>
			<td>12566014</td>
			<td>Reagent</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">Syringe Filter</td>
			<td>EMD Millipore</td>
			<td>SLHV033RS</td>
			<td>0.45&#160;&#181;m&#160;</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">HiTrap heparin columns</td>
			<td>gelifesciences</td>
			<td>17-0406-01</td>
			<td>1 mL&#160;</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">Amicon ultra-4 centrifugal filter</td>
			<td>EMD Millipore</td>
			<td>&#160;UFC810024</td>
			<td>100,000 MWCO</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">Benzonase nuclease</td>
			<td>Sigma-Aldrich</td>
			<td>E1014</td>
			<td>Reagent</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">Sodium deoxycholate</td>
			<td>Sigma-Aldrich</td>
			<td>D5670</td>
			<td>Make fresh solution for each batch</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">mouse stereotaxic apparatus</td>
			<td>B&#38;E TEKSYSTEMS LTD</td>
			<td>#SR-5M/6M</td>
			<td>Equipment</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">pentobarbital</td>
			<td>SigmaAldrich</td>
			<td>#1507002</td>
			<td>Reagent</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">mouse stereotaxic apparatus</td>
			<td>B&#38;E TEKSYSTEMS LTD</td>
			<td>#SR-5M/6M</td>
			<td>Equipment</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">Hydrogen peroxide solution</td>
			<td>SigmaAldrich</td>
			<td>#216763</td>
			<td>Reagent</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">Optical Fiber</td>
			<td>Thorlabs</td>
			<td>FT200EMT</td>
			<td>0.39 NA, &#216;200 &#181;m</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">microsyringe pump</td>
			<td>Nanoliter 2000 Injector, WPI</td>
			<td></td>
			<td>Equipment</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">ceramic ferrule</td>
			<td>Shanghai Fiblaser</td>
			<td></td>
			<td>230&#8201;&#956;m I.D., 2.5 mm O.D.</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">Gene Observer</td>
			<td>BiolinkOptics</td>
			<td></td>
			<td>Equipment</td>
		</tr>
	</tbody>
</table>

in vivo monitoring of circadian clock gene expression in the mouse suprachiasmatic nucleus using fluorescence reporters

This technique combines optical fiber mediated fluorescence recordings with the precise delivery of recombinant adeno-associated virus based gene reporters. This new and easy to use in vivo fluorescence monitoring system was developed to record the transcriptional rhythm of the clock gene, Cry1, in the suprachiasmatic nucleus (SCN) of freely moving mice. To do so, a Cry1 transcription fluorescence reporter was designed and packaged into Adeno-associated virus. Purified, concentrated virus was injected into the mouse SCN followed by the insertion of an optic fiber, which was then fixed onto the surface of the brain. The animals were returned to their home cages and allowed a 1-month post-operative recovery period to ensure sufficient reporter expression. Fluorescence was then recorded in freely moving mice via an in vivo monitoring system that was constructed at our institution. For the in vivo recording system, a 488 nm laser was coupled with a 1 &#215; 4 beam-splitter that divided the light into four laser excitation outputs of equal power. This setup enabled us to record from four animals simultaneously. Each of the emitted fluorescence signals was collected via a photomultiplier tube and a data acquisition card. In contrast to the previous bioluminescence in vivo circadian clock recording technique, this fluorescence in vivo recording system allowed the recording of circadian clock gene expression during the light cycle.

Fluorescence reporter design of Cry1 was show in Figure 1A. Using the approach detailed above, 500 nL of rAAV-P(Cry1)-intron336-Venus-NLS-D2 was successfully injected into the SCN of an adult mouse, and exhibited robust Venus expression (Figure 1B, 1C). Fluorescence signals recorded under 12h/12h light/dark (LD) and dark/dark (DD) conditions (Figure 2) yielded robust circadian rhythm in both conditions. Finally, fluorescence signal was recorded in long and short photoperiod conditions (Figure 3).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56765/56765fig1.jpg" /> 
Figure 1. Fluorescence reporter design and it expression in the SCN 
(A) The fluorescence reporter design of Cry1. (B) Photomicrograph of the mouse brain showing reporter expression 8 days after the virus injection. (C) Magnification of the area boxed in red on the photomicrograph shown in (B). <a href="/files/ftp_upload/56765/56765fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56765/56765fig2.jpg" /> 
Figure 2. Fluorescence in vivo recording in the SCN under LD and DD conditions 
(A) Background fluorescence in the SCN showing no detectable rhythm under both LD and DD conditions. The white and gray areas indicate light-on and light-off periods, respectively. (B) P(Cry1)-intron336-Venus-NLS-D2 expression over the course of 7 days in a 12-12 LD condition and 7 days in a DD condition. (C) Analysis of circadian rhythm in the LD condition (~24.4 h); the red line indicates the converged fit in the Sine curve (the result of the fit is shown in the left panel). (D) Analysis of circadian rhythm in the DD condition (~23.25 h); the red line indicates the converged fit in the Sine curve (the result of the fit is shown in the left panel). <a href="/files/ftp_upload/56765/56765fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56765/56765fig3.jpg" /> 
Figure 3. In vivo fluorescence recordings in the SCN under 20-4 long and 4-20 short photoperiod conditions 
Example of a fluorescence recording of P(Cry1)-intron336-Venus-NLS-D2 expression in the SCN for 7 days under a 12-12 LD condition, followed by 7 days under a 20-4 long photoperiod condition, 3 days under a DD condition, 3 days under a 12-12 LD condition, and then 7 days under a 4-20 short photoperiod condition. White and gray areas indicate the light-on and light-off periods, respectively. <a href="/files/ftp_upload/56765/56765fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1. Specific reagents required for protocol (See Table of Materials)
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>primer name</td>
			<td colspan="3">sequence</td>
		</tr>
		<tr>
			<td>primer F1</td>
			<td colspan="3">cactaggggttcctgcggccgcACGCGTGTAAAGATGCACATGTG</td>
		</tr>
		<tr>
			<td>primer R1</td>
			<td colspan="3">TTGTAACCTTGATACTTACCTACTTAGATCGCAGATCTCGTCCGG</td>
		</tr>
		<tr>
			<td>primer F2</td>
			<td colspan="3">GTTATGACACAGTGTAGAAACTATGGCATAGGACAGATGACTGTG</td>
		</tr>
		<tr>
			<td>primer R2</td>
			<td colspan="3">CATGGTCTTTGTAGTCCATGGTGGGTACCtCTTGACAGCTCTACC</td>
		</tr>
		<tr>
			<td>primer F3</td>
			<td colspan="3">ctgtattttcagggcCCTGCAGGtGTGAGCAAGGGCGAGGAGCTG</td>
		</tr>
		<tr>
			<td>primer R3</td>
			<td colspan="3">TACCTTTCTCTTCTTTTTTGGAGGCTTGTACAGCTCGTCCATGCC</td>
		</tr>
		<tr>
			<td>primer F4</td>
			<td colspan="3">GCATGGACGAGCTGTACAAGCCTCCAAAAAAGAAGAGAAAGGTAG</td>
		</tr>
		<tr>
			<td>primer R4</td>
			<td colspan="3">tgatatcgaattcGGATCCCTACACATTGATCCTAGCAGAAGCAC</td>
		</tr>
	</tbody>
</table>
Table 2. Primer used in the protocol
Table 3. DNA sequence used in the protocol <a href="/files/ftp_upload/56765/Table3.xlsx">Please click here to download this file.</a>

Watch this Scientific Journal Video about In Vivo Monitoring of Circadian Clock Gene Expression in the Mouse Suprachiasmatic Nucleus Using Fluorescence Reporters at JoVE.com

In Vivo Monitoring of Circadian Clock Gene Expression in the Mouse Suprachiasmatic Nucleus Using Fluorescence Reporters

This newly developed fluorescence-based technology enables long-term monitoring of the transcription of circadian clock genes in the suprachiasmatic nucleus (SCN) of freely moving mice in real-time and at a high temporal resolution.

In Vivo Monitoring of Circadian Clock Gene Expression in the Mouse Suprachiasmatic Nucleus Using Fluorescence Reporters

This technique combines optical fiber mediated fluorescence recordings with the precise delivery of recombinant adeno-associated virus based gene ...

in-vivo-monitoring-circadian-clock-gene-expression-mouse

Nanyang Technological University

Research

JoVE Journal

Neuroscience

8.2K Views.  Peking University. This newly developed fluorescence-based technology enables long-term monitoring of the transcription of circadian clock genes in the suprachiasmatic nucleus (SCN) of freely moving mice in real-time and at a high temporal resolution.

Video: In Vivo Monitoring of Circadian Clock Gene Expression in the Mouse Suprachiasmatic Nucleus Using Fluorescence Reporters

Slice Preparation, Organotypic Tissue Culturing and Luciferase Recording of Clock Gene Activity in the Suprachiasmatic Nucleus

A central circadian (~24 hr) clock coordinating daily rhythms in physiology and behavior resides in the suprachiasmatic nucleus (SCN) located in the anterior hypothalamus. The clock is directly synchronized by light via the retina and optic nerve. Circadian oscillations are generated by interacting negative feedback loops of a number of so called "clock genes" and their protein products, including the Period (Per) genes. The core clock is also dependent on membrane depolarization, calcium and cAMP 1. The SCN shows daily oscillations in clock gene expression, metabolic activity and spontaneous electrical activity. Remarkably, this endogenous cyclic activity persists in adult tissue slices of the SCN 2-4. In this way, the biological clock can easily be studied in vitro, allowing molecular, electrophysiological and metabolic investigations of the pacemaker function.
The SCN is a small, well-defined bilateral structure located right above the optic chiasm 5. In the rat it contains ~8.000 neurons in each nucleus and has dimensions of approximately 947 &mu;m (length, rostrocaudal axis) x 424 &mu;m (width) x 390 &mu;m (height) 6. To dissect out the SCN it is necessary to cut a brain slice at the specific level of the brain where the SCN can be identified. Here, we describe the dissecting and slicing procedure of the SCN, which is similar for mouse and rat brains. Further, we show how to culture the dissected tissue organotypically on a membrane 7, a technique developed for SCN tissue culture by Yamazaki et al. 8. Finally, we demonstrate how transgenic tissue can be used for measuring expression of clock genes/proteins using dynamic luciferase reporter technology, a method that originally was used for circadian measurements by Geusz et al. 9. We here use SCN tissues from the transgenic knock-in PERIOD2::LUCIFERASE mice produced by Yoo et al. 10. The mice contain a fusion protein of PERIOD (PER) 2 and the firefly enzyme LUCIFERASE. When PER2 is translated in the presence of the substrate for luciferase, i.e. luciferin, the PER2 expression can be monitored as bioluminescence when luciferase catalyzes the oxidation of luciferin. The number of emitted photons positively correlates to the amount of produced PER2 protein, and the bioluminescence rhythms match the PER2 protein rhythm in vivo 10. In this way the cyclic variation in PER2 expression can be continuously monitored real time during many days. The protocol we follow for tissue culturing and real-time bioluminescence recording has been thoroughly described by Yamazaki and Takahashi 11.

The procedure of preparing slices containing the adult mouse hypothalamic suprachiasmatic nucleus (SCN), and a rapid way to culture the SCN tissue in organotypic culture condition, are reported. Further, the measurement of oscillatory clock gene protein expression using dynamic luciferase reporter technology is described.

A central circadian (~24 hr) clock coordinating daily rhythms in physiology and behavior resides in the suprachiasmatic nucleus (SCN) located in the ...

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Non-Laser Capture Microscopy Approach for the Microdissection of Discrete Mouse Brain Regions for Total RNA Isolation and Downstream Next-Generation Sequencing and Gene Expression Profiling

As technological platforms, approaches such as next-generation sequencing, microarray, and qRT-PCR have great promise for expanding our understanding of the breadth of molecular regulation. Newer approaches such as high-resolution RNA sequencing (RNA-Seq)1 provides new and expansive information about tissue- or state-specific expression such as relative transcript levels, alternative splicing, and micro RNAs2-4. Prospects for employing the RNA-Seq method in comparative whole transcriptome profiling5 within discrete tissues or between phenotypically distinct groups of individuals affords new avenues for elucidating molecular mechanisms involved in both normal and abnormal physiological states. Recently, whole transcriptome profiling has been performed on human brain tissue, identifying gene expression differences associated with disease progression6. However, the use of next-generation sequencing has yet to be more widely integrated into mammalian studies.
Gene expression studies in mouse models have reported distinct profiles within various brain nuclei using laser capture microscopy (LCM) for sample excision7,8. While LCM affords sample collection with single-cell and discrete brain region precision, the relatively low total RNA yields from the LCM approach can be prohibitive to RNA-Seq and other profiling approaches in mouse brain tissues and may require sub-optimal sample amplification steps. Here, a protocol is presented for microdissection and total RNA extraction from discrete mouse brain regions. Set-diameter tissue corers are used to isolate 13 tissues from 750-&mu;m serial coronal sections of an individual mouse brain. Tissue micropunch samples are immediately frozen and archived. Total RNA is obtained from the samples using magnetic bead-enabled total RNA isolation technology. Resulting RNA samples have adequate yield and quality for use in downstream expression profiling. This microdissection strategy provides a viable option to existing sample collection strategies for obtaining total RNA from discrete brain regions, opening possibilities for new gene expression discoveries.

Non-Laser Capture Microscopy Approach for the Microdissection of Discrete Mouse Brain Regions for Total RNA Isolation and Downstream Next-Generation Sequencing and Gene Expression Profiling  |

RNA expression profiling of discrete mouse brain regions requires a precise and repeatable tissue collection strategy. A protocol that uses both coronal brain sectioning and tissue corer-assisted microdissection is described here. The yield and quality of total RNA obtained from the resulting samples confirms the utility of the outlined method.

As technological platforms, approaches such as next-generation sequencing, microarray, and qRT-PCR have great promise for expanding our understanding of ...

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central nervous system (CNS). Although much is known about the molecular mechanisms that pattern the spinal cord and elicit neuronal differentiation1, 2, we lack a deep understanding of these early events at the level of cell behavior. It is thus critical to study the behavior of neural progenitors in real time as they undergo neurogenesis.
In the past, real-time imaging of early embryonic tissue has been limited by cell/tissue viability in culture as well as the phototoxic effects of fluorescent imaging. Here we present a novel assay for imaging such tissue for long periods of time, utilizing a novel ex vivo slice culture protocol and wide-field fluorescence microscopy (Fig. 1). This approach achieves long-term time-lapse monitoring of chick embryonic spinal cord progenitor cells with high spatial and temporal resolution.
This assay may be modified to image a range of embryonic tissues3, 4 In addition to the observation of cellular and sub-cellular behaviors, the development of novel and highly sensitive reporters for gene activity (for example, Notch signaling5) makes this assay a powerful tool with which to understand how signaling regulates cell behavior during embryonic development.

Imaging embryonic tissue in real-time is challenging over long periods of time. Here we present an assay for monitoring cellular and sub-cellular changes in chick spinal cord for long periods with high spatial and temporal resolution. This technique can be adapted for other regions of the nervous system and developing embryo.

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

Micromanipulation of Gene Expression in the Adult Zebrafish Brain Using Cerebroventricular Microinjection of Morpholino Oligonucleotides

Manipulation of gene expression in tissues is required to perform functional studies. In this paper, we demonstrate the cerebroventricular microinjection (CVMI) technique as a means to modulate gene expression in the adult zebrafish brain. By using CVMI, substances can be administered into the cerebroventricular fluid and be thoroughly distributed along the rostrocaudal axis of the brain. We particularly focus on the use of antisense morpholino oligonucleotides, which are potent tools for knocking down gene expression in vivo. In our method, when applied, morpholino molecules are taken up by the cells lining the ventricular surface. These cells include the radial glial cells, which act as neurogenic progenitors. Therefore, knocking down gene expression in the radial glial cells is of utmost importance to analyze the widespread neurogenesis response in zebrafish, and also would provide insight into how vertebrates could sustain adult neurogenesis response. Such an understanding would also help the efforts for clinical applications in human neurodegenerative disorders and central nervous system regeneration. Thus, we present the cerebroventricular microinjection method as a quick and efficient way to alter gene expression and neurogenesis response in the adult zebrafish forebrain. We also provide troubleshooting tips and other useful information on how to carry out the CVMI procedure.

In this article, we demonstrate a method for manipulation of gene expression in the ventricular cells of the adult zebrafish telencephalon using antisense morpholino oligonucleotides. We present this method as an efficient and quick protocol that can be used for functional studies in the adult vertebrate brain.

Manipulation of gene expression in tissues is  required to perform functional studies. In this paper, we demonstrate the  cerebroventricular ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Analysis of Gene Expression Changes in the Rat Hippocampus After Deep Brain Stimulation of the Anterior Thalamic Nucleus

Deep brain stimulation (DBS) surgery, targeting various regions of the brain such as the basal ganglia, thalamus, and subthalamic regions, is an effective treatment for several movement disorders that have failed to respond to medication. Recent progress in the field of DBS surgery has begun to extend the application of this surgical technique to other conditions as diverse as morbid obesity, depression and obsessive compulsive disorder. Despite these expanding indications, little is known about the underlying physiological mechanisms that facilitate the beneficial effects of DBS surgery. One approach to this question is to perform gene expression analysis in neurons that receive the electrical stimulation. Previous studies have shown that neurogenesis in the rat dentate gyrus is elicited in DBS targeting of the anterior nucleus of the thalamus1. DBS surgery targeting the ATN is used widely for treatment refractory epilepsy. It is thus of much interest for us to explore the transcriptional changes induced by electrically stimulating the ATN. In this manuscript, we describe our methodologies for stereotactically-guided DBS surgery targeting the ATN in adult male Wistar rats. We also discuss the subsequent steps for tissue dissection, RNA isolation, cDNA preparation and quantitative RT-PCR for measuring gene expression changes. This method could be applied and modified for stimulating the basal ganglia and other regions of the brain commonly clinically targeted. The gene expression study described here assumes a candidate target gene approach for discovering molecular players that could be directing the mechanism for DBS.

The mechanism underlying the therapeutic effects of Deep Brain Stimulation (DBS) surgery needs investigation. The methods presented in this manuscript describe an experimental approach to examine the cellular events triggered by DBS by analyzing the gene expression profile of candidate genes that can facilitate neurogenesis post DBS surgery.

Deep brain stimulation (DBS) surgery, targeting various regions of the brain such as the basal ganglia, thalamus, and subthalamic regions, is an effective ...