We thank Dr. Maiken Nedergaard (University of Rochester Medical Center) for the head plate design. The work has been supported by NIH awards to S.D (R01DA026325 and P30AI078498 and foundation grants to KK (DANA foundation Brain and Immunoimaging program, American Heart Association 0635595T and the ALS Association [#1112)]).

1. Preparing the animal for imaging
Note: We use adult c57BL mice. Different transgenic strains can be used depending on the research question. Microvascular surgeries and pulse oximetry are facilitated in mice with white fur (e.g. FVB strain).
<ol>
	<li>Prepare the surgical site at the bench. All surgical instruments need to be autoclaved or disinfected with 70% ethanol.</li>
	<li>Insert a rectal probe, and continuously measure the animal's body temperature throughout the procedure</li>
	<li>Anesthetize the mouse using initially 5% isoflurane. In 15-30 s, when the animal is not moving, place it on a heating pad (37 &deg;C), and apply a face mask for the continuous delivery of air containing 1.3-1.5% isoflurane). Make sure that the animal is not responding to a pain stimulus (e.g. tail pinch).</li>
	<li>Use artificial tear gel to cover the mouse's eyes, to prevent exposure to dry air.</li>
	<li>Using an electric razor, remove hair from the head and from both thighs. Apply hair remover (e.g. Nair) to the thigh for 2 min, and then wipe it carefully to remove the remaining hair. This thigh will be used for oximetry.</li>
	<li>Disinfect the scalp using a 10% povidone-iodine solution and 70% ethanol.</li>
</ol>
2. Preparing the open skull cranial window
<ol>
	<li>Starting 5 mm caudal to the skull, make an incision in the scalp with scissors, and advance one centimeter. Move skin to the sides to expose the skull.</li>
	<li>To remove the membranes on top of the skull apply 10% ferric chloride solution to the top of the skull. Wipe away the excess solution, and scrape the membranes away with 45&deg; angle #5 tweezers. It is important to prepare the site carefully, in order to ensure that the head plate can be securely attached to the skull (steps #4 thru 6).</li>
	<li>Using a dissection microscope, identify the area of interest. Note that the cranial sutures should be avoided because the skull can break unpredictably along these lines.</li>
	<li>Apply a thin layer of rapid glue (e.g. Locktite 454) around the edges of the window of the head plate (Figure 1). Position the head plate in such a way that the area of interest is exposed in the window. Apply light pressure on the head plate. Apply a small amount of dental cement to polymerize the glue rapidly. Hold for 10 seconds while the glue is polymerizing.</li>
	<li>Apply a small amount of rapid glue to the edges of the window to seal the head plate and the skull. Screw the head plate to the animal holder under the dissection scope.</li>
	<li>Using the Microtorque II drill set at 6000 rpm and a IRF 005 drill bit, remove all extra glue from the skull and from the head plate. Any remaining glue on the head plate should be removed, since it will otherwise interfere with the attachment of the glass cover.</li>
	<li>Set the drill at 1000 rpm, and start drilling the skull by making a circle inside the head plate window, diameter ~5 mm. Use light sweeping movements as if you are drawing with a very soft pencil. Stop drilling every 20-30 seconds to remove bone dust using compressed air. 
	Note: The shape of the opening in the head plate allows additional access for the drill bit (for left- and right-handed surgeon), which makes it easier to create a circular cranial window. In addition, the two odd shaped holes provide the required space to insert a Clark-electrode or glass-electrode into the brain tissue under the cranial window for additional polarographic or electrophysiological measurements.</li>
	<li>As the drilling progresses, a burrow is formed around the intact skull in the center. Be careful not to poke through the thinned skull, but to make this burrow of even thickness. Be extra careful around large blood vessels -they should not be compromised and, if possible, not even touched.</li>
	<li>Use one "leg" of tweezers #3 to lift ("flick off") the bone in the center of the window. Apply a drop of artificial cerebrospinal fluid (aCSF) on the window.</li>
	<li>Wipe aCSF gently using a corner of soft tissue paper (e.g. Kimwipe). Usually the dura is damaged in one or more places around the edge of the window. Starting at one of these sites, gently lift and then remove the dura from the brain surface, using 45&deg; angle tweezers #5. When choosing the site at which to start removing the dura, avoid the areas adjacent to large blood vessels. If bleeding occurs, apply a drop of aCSF and wait for 2-3 min for minor bleeding to stop.</li>
	<li>Prepare 0.07% low-melting agarose dissolved in aCSF) Bring the melted agarose to body temperature. Wipe away the excess of aCSF from the surface of the brain. Pour the agarose in the window, and cover with a glass slide. Press the glass gently to make a contact between the glass and the head plate, and wait 10-20 sec until the agarose is solid.</li>
	<li>Remove excess agarose from the glass cover using tweezers and soft tissue paper.</li>
	<li>Put a small amount of rapid glue around the glass to glue the glass to the head plate. Apply a small amount of dental cement to solidify the glue (Figure 1).</li>
</ol>
3. Intravenous injection and blood oxygenation monitoring
Note: We routinely used the femoral vein for intravenous application of Texas red rather than the tail vein. This is because femoral vein injections provide safe and reproducible bolus injections of the dye.
<ol>
	<li>Turn the animal partially over to expose the inside of the left thigh. Use tape to secure the animal in this position.</li>
	<li>Apply antiseptic scrub, and then 100 % ethanol to the skin.</li>
	<li>Make an incision along the medial thigh from the knee to the pubic symphysis. Separate soft tissues by blunt dissection using tweezers #5.</li>
	<li>Fill 1ml syringe with 130 &mu;l of Texas Red (2.5 mg/ml). Attach a new needle (gauge 30), and bend it carefully at 30&deg;, keeping the bevel up. Fill the needle with Texas Red solution.</li>
	<li>Under a dissection microscope, insert the needle into the femoral vein and inject 100 &mu;l of Texas Red. Remove needle and lightly press the vein with gauze to stop bleeding.</li>
	<li>Close the skin using 4-0 suture.</li>
	<li>For continuous monitoring of blood SpO2 (to ensure adequate blood oxygenation) place the oximeter probe on the right thigh (from which hair was previously removed).</li>
</ol>
4. Two-photon imaging
Note: We use an Olympus Fluoview1000 multiphoton imaging system (upright) with a Spectra-Physics MaiTai HP DeepSee femtosecond Ti:Sa laser as an excitation source. For imaging, we use &times; 10 NA 0.45 (Zeiss C-Apochromat), or &times; 25 NA 1.05 (Olympus XLPlan N) water-immersion objectives. We take images at 12-bit depth at a resolution of 512 &times; 512 pixels with a pixel dwell time of 2 &mu;s. The NADH and Texas-Red-dextran are concurrently excited at 740 nm, and fluorescence is separated from the excitation light using a dichroic mirror/near-IR-blocking filter combination (FF665-Di01; FF01-680/SP, Semrock, Rochester, NY, USA) divided into two channels using a dichroic mirror (505DCXRU, Chroma, Rockingham, VT, USA), and bandpass filtered (NADH-Semrock FF460/80Texas-Red-dextran-Semrock FF607/36). Laser power measured after the objective was 10-20 mW for imaging in layer I and 20-50 mW for imaging in layer II.
<ol>
	<li>Transfer the surgically prepared mouse to the microscope stage. Take an initial picture of the cranial window site using bright field illumination at 4x magnification to use as a reference map for registration with higher magnification two-photon angiographies.</li>
	<li>Zoom in on the area of interest using &times; 10 magnification. Select an area of interest in cortical layers I or II (up to 150 &mu;m below the pial surface). If higher magnification is desired, use &times; 25 objective</li>
	<li>Start record the time series (Figure 2). We recommend using an average of 3-5 frames to increase the quality of the image.</li>
	<li>Induce hypoxemia by adding 50 % N2 to the air that the animal is breathing. This will bring the O2 level to 10 %. Verify hypoxemia by monitoring oximeter data.</li>
	<li>Continue to record the time series through hypoxemia (for example, for 30 seconds), and after normal O2 level is restored.</li>
	<li>Collect data for calculation of the oxygen distribution by detecting, measuring and quantifying the area of hypoxia and oxygenated tissue cylinders around the blood vessels.</li>
</ol>
5. Data processing
<ol>
	<li>Determine the Krogh tissue cylinder radius R. This can be done manually (Figure 3) by measuring the distance between center of the blood vessel and the boundary at which high NADH fluorescence is detected.</li>
	<li>Alternatively, use the computational investigator-independent semi-automated determination of the tissue cylinder radius R and the central blood vessel r that has been described previously1 in supplementary form and summarized in the discussion section of this protocol.</li>
	<li>Determine the area of hypoxia using open-access image analysis software, e.g. ImageJ. To do this, use NADH signal, define a threshold that delineates the high NADH intensity area, and then measure the area with elevated NADH tissue fluorescence.</li>
</ol>
6. Representative Results
Figure 2 Shows a video of NADH/angiography images during transient hypoxemia (for the PDF version of this Figure, selected still images have been used). The inspired oxygen concentration was lowered from 21 % to 10 % for a period of 3 min. This level of experimentally induced hypoxemia is sufficient to induce severe hypoxia in the cerebral cortex. Hypoxia led to an increase in NADH fluorescence, initially in the areas which were furthest from the arterial blood supply. Note the sharp NADH tissue boundaries, which represent observable tissue boundaries of oxygen diffusion from the cortical microcirculation. The image in Figure 3 shows the geometry of oxygen diffusion boundaries surrounding cerebral vessels. It is possible to deduce Krogh cylinder diameters from these boundaries, as described previously 1.
<img alt="Figure 1" src="/files/ftp_upload/3466/3466fig1.jpg" /> 
Figure 1. (A) Mouse with cranial window prepared for imaging. (B) Dimensions of head plate.
<img alt="Figure 2" src="/files/ftp_upload/3466/3466fig2.jpg" /> 
Figure 2. Endogenous NADH green fluorescence visualized with two-photon imaging through cranial window, approximately 50 &mu;m below the pial surface. Blood vessels are visualized with Texas Red dextran. Oxygen in inhaled air was reduced to 10% for 3 min, and then restored for 21%. Scale bar: 50&mu;m.
<img alt="Figure 3" src="/files/ftp_upload/3466/3466fig3.jpg" /> 
Figure 3. Geometry of NADH fluorescence boundaries. Yellow outline shows boundary of functional hypoxia; blue circles show projected oxygenated (Krogh) cylinders. Blue lines show cylinder radii; numbers show radii in micrometers.

<ol>
	<li>Kasischke, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+NADH+imaging+exposes+boundaries+of+oxygen+diffusion+in+cortical+vascular+supply+regions.">Two-photon NADH imaging exposes boundaries of oxygen diffusion in cortical vascular supply regions.</a> J. Cereb. Blood. Flow. Metab. 31, 68-81 (2011).</li><li>Ndubuizu, O., LaManna, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brain+tissue+oxygen+concentration+measurements.">Brain tissue oxygen concentration measurements.</a> Antioxid. Redox. Signal. 9, 1207-1219 (2007).</li><li>Sharan, M., Vovenko, E. P., Vadapalli, A., Popel, A. S., Pittman, R. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+and+theoretical+studies+of+oxygen+gradients+in+rat+pial+microvessels.">Experimental and theoretical studies of oxygen gradients in rat pial microvessels.</a> J. Cereb. Blood. Flow. Metab. 28, 1597-1604 (2008).</li><li>Sakadzic, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+high-resolution+measurement+of+partial+pressure+of+oxygen+in+cerebral+vasculature+and+tissue.">Two-photon high-resolution measurement of partial pressure of oxygen in cerebral vasculature and tissue.</a> Nat. Methods. 7, 755-759 (2010).</li><li>Vishwasrao, H. D., Heikal, A. A., Kasischke, K. A., Webb, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conformational+dependence+of+intracellular+NADH+on+metabolic+state+revealed+by+associated+fluorescence+anisotropy.">Conformational dependence of intracellular NADH on metabolic state revealed by associated fluorescence anisotropy.</a> J. Biol. Chem. 280, 25119-25126 (2005).</li><li>Kleinfeld, D., Mitra, P. P., Helmchen, F., Denk, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluctuations+and+stimulus-induced+changes+in+blood+flow+observed+in+individual+capillaries+in+layers+2+through+4+of+rat+neocortex.">Fluctuations and stimulus-induced changes in blood flow observed in individual capillaries in layers 2 through 4 of rat neocortex.</a> Proc. Natl. Acad. Sci. U. S. A. 95, 15741-15746 (1998).</li><li>Brown, W. R., Thore, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Review:+cerebral+microvascular+pathology+in+ageing+and+neurodegeneration.">Review: cerebral microvascular pathology in ageing and neurodegeneration.</a> Neuropathol. Appl. Neurobiol. 37, 56-74 (2011).</li><li>de la Torre, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impaired+brain+microcirculation+may+trigger+Alzheimer's+disease.">Impaired brain microcirculation may trigger Alzheimer's disease.</a> Neurosci. Biobehav. Rev. 18, 397-401 (1994).</li><li>Brown, C. E., Boyd, J. D., Murphy, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Longitudinal+in+vivo+imaging+reveals+balanced+and+branch-specific+remodeling+of+mature+cortical+pyramidal+dendritic+arbors+after+stroke.">Longitudinal in vivo imaging reveals balanced and branch-specific remodeling of mature cortical pyramidal dendritic arbors after stroke.</a> J. Cereb. Blood. Flow. Metab. 30, 783-791 (2010).</li><li>Wilson, D. F., Erecinska, M., Drown, C., Silver, I. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+oxygen+dependence+of+cellular+energy+metabolism.">The oxygen dependence of cellular energy metabolism.</a> Arch. Biochem. Biophys. 195, 485-493 (1979).</li></ol>

The authors have nothing to disclose.

High spatial resolution information about oxygen diffusion is important to understanding how blood flow in the brain is regulated to provide oxygen to brain cells, and to meet metabolic demand. Traditional polarographic oxygen measurements using Clark-style glass electrodes are highly invasive and have low spatial resolution2-3 and significant (second range) response time. So far the only non-invasive method for measuring pO2 in brain tissue is phosphorescence quenching, where the rate of decay of the excited probe is proportional to oxygen concentration4. This method provides accurate oxygen concentrations, but requires a proprietary dye and a technically sophisticated phosphorescence lifetime imaging system. Here, we demonstrate a straightforward, simpler approach that can be conducted on a standard two-photon imaging system with two flurescence channels. Our approach takes advantage of an intrinsic tissue signal5 and only requires contrasting visualization of the cortical microcirculation. Because of the non-linear, essentially binary increase of NADH fluorescence at functionally limiting oxygen concentrations1, increased intrinsic NADH fluorescence is observed only in areas with significant, metabolically limiting hypoxia. An important implication is that tissue boundaries of oxygen diffusion from cortical microvessels are directly observable by cylindrically shaped intensity changes of endogenous NADH fluorescence. We refer to these structures as Krogh cylinders, because the concept of cylindrically shaped structures that define the oxygenated volume of tissue surrounding a blood vessel was introduced by August Krogh and has been recently experimentally observed using two-photon NADH imaging1. Krogh cylinders images can be collected in 3D by taking a z-stack of image frames. They are especially prominent in the vicinity of penetrating arterioles and they are congruous with capillary depleted periarteriolar tissue cylinders1,4. 
To provide an objective determination of the Krogh tissue cylinder radius R (see Section 5.2) we measured their radial pixel intensity values within a well-defined segment between the center of the cylinder and the outer boundary using the Matlab function &#x201C;improfile&#x201D;. The outer boundary of the segment should be chosen to extend with a safety margin beyond the visible boundary. To improve the signal-to-noise level we averageed over all radial lines needed to cover the visible cylinder segment at 1&deg; steps. The resulting mean radial intensity profile within the segment exhibited a steep increase which corresponded to the visible tissue boundary R. The we fit a sigmoidal function (e.g. Boltzmann function) to the averaged radial intensity profile and used its inflection point (also known as x0) as a definition of R. The corresponding two-photon microangiography (Texas-red) revealed the cross-section of a solitary central blood vessel in the center of cylinder. The diameter of the central blood vessel can be directly applied to determine r.
Two-photon NADH imaging provides the same spatial resolution as the concurrent high-resolution imaging of the cortical microangiography. An important feature for the quantitative application of this method is that p50 of the NADH fluorescence increase has been measured to be of 3.4 &plusmn; 0.6&#x2009;mm&#x2009;Hg 1 and that the NADH fluorescence intensity as a function of microregional tissue pO2 can be mathematically described with a sigmoidal function. . We show that this technique allows one to identify brain areas which are most vulnerable to hypoxia (by decreasing oxygen content in the air to 10 %). We also show that oxygen diffusion follows a simple geometric perivascular pattern. 
One critical step for this method is the quality of the cranial window preparation. The surgery should produce minimal damage in order not to disturb blood flow to the exposed area. A concern is that in a surgically compromised preparation, the cortex under the window may be hypoxic to begin with, precluding any meaningful experiments. A well-prepared cranial window should have intact major and minor blood vessels with vivid blood flow in all vessel types and no significant bleeding along the edges. Under normoxic conditions (PaO2 80-100 mmHg, Sp O2 97-99%) the brain parenchyma should exhibit uniform, homogeneous NADH fluorescence without conspicuous, bright tissue patches with elevated NADH fluorescence. 
A fundamental physical constraint of our approach is limited depth penetration. The blue-green NADH fluorescence in brain is rapidly attenuated by hemoglobin absorption and tissue scattering at these wavelengths. Even with high numerical aperture (e.g. 1.05) water immersion objectives two-photon NADH imaging is currently limited to cortical layers I and II. This limitation is scientifically relevant because energy metabolism in or in proximity to white matter will likely differ from gray matter. However, the investigation of deep cortical structures such as layers IV-VI or subcortical structures such as white matter tracts or the striatum would require the use of specialized microlenses as described in the mouse cortex in vivo6.
NADH-based measurement of oxygen diffusion boundaries can be especially useful when combined with other measurements such as analyses of functional hyperemia, and detection of capillary flux rates7. For example, this technique can be adapted&nbsp;to visualize hypoxia in stroke and Alzheimer&#x2019;s disease (AD) models. The simple geometry of oxygen diffusion allows one to predict the oxygen gradient in microvascular beds under circumstances where capillary density is decreased8 (e.g., AD9) and to examine whether brain tissue regions with reduced capillary density are at increased risk for hypoxia damage due to microstrokes. The ability to image microregionally also allows one to examine the geometry and size of tissue microstrokes and determine the volume of tissue in which hypoxia occurs, as well as the relationship between tissue hypoxia and subsequent neuronal death or capillary remodeling10. 
Finally, since increases in endogenous NADH fluorescence are the direct consequence of acute mitochondrial dysfunction, this method creates the opportunity to use NADH imaging as a specific reporter for neural energy metabolism11 and a proxy for mitochondrial dysfunction. 
In conclusion, two-photon imaging of endogenous NADH fluorescence is a simple, undemanding tool that can be used to understand oxygen delivery and consumption in the brain under both normal and in pathologic states.

<table border="1">
<tbody>
 <tr valign="top">
 <td>Name 			of the reagent</td>
 <td>Company</td>
 <td>Catalogue 			number</td>
 <td>Comments 			(optional)</td>
 </tr>
 <tr>
 <td>Heating 			pads</td>
 <td>Beyond 			Bodi Heat</td>
 <td>&nbsp;</td>
 <td>&nbsp; </td>
 </tr>
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 <td>Ophthalmic 			ointment (Artificial tears)</td>
 <td>Pfizer&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
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 <tr>
 <td>Povidone-iodine 			10% solution</td>
 <td>Betadine</td>
 <td>&nbsp;</td>
 <td>&nbsp; </td>
 </tr>
 <tr>
 <td>Ferric 			chloride 10% solution</td>
 <td>&nbsp; </td> 
 <td>&nbsp; </td>
 <td>&nbsp; </td>
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 <td>Cement</td>
 <td>Stoelting 			Company</td>
 <td>&nbsp;51456</td>
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 </td>
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 <td>Cyanoacrylate 			454</td>
 <td>Loctite</td>
 <td>&nbsp;</td>
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 </td>
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 <td>aCSF</td>
 <td>Harvard 			Apparatus</td>
 <td>597316</td>
 <td>&nbsp; </td>
 </tr>
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 <td>Microtorque 			II handpiece kit</td>
 <td>Pearson</td>
 <td>R14-0002</td>
 <td>&nbsp; </td>
 </tr>
 <tr>
 <td>IRF 			007 drill bits</td>
 <td>Fine 			Science Tools</td>
 <td>19008-07</td>
 <td>&nbsp; </td>
 </tr>
 <tr>
 <td>Forceps 			#5</td>
 <td>Fine 			Science Tools</td>
 <td>11295</td>
 <td>&nbsp; </td>
 </tr>
 <tr>
 <td>Forceps 			#5/45</td>
 <td>Fine 			Science Tools</td>
 <td>11251-35</td>
 <td>&nbsp; </td>
 </tr>
 <tr>
 <td>#0 			glass coverslip</td>
 <td>Electron 			Microscopy Sciences</td>
 <td>63750-01</td>
 <td>&nbsp; </td>
 </tr>
 <tr>
 <td>Photomultiplier 			tube</td>
 <td>Hamamatsu</td>
 <td>HC125-02</td>
 <td>&nbsp; </td>
 </tr>
 <tr>
 <td lang="it-IT" xml:lang="it-IT">Ti:Sapphire 			laser Mai-Tai</td>
 <td>Spectra-Physics</td>
 <td>&nbsp;</td>
 <td>&nbsp; </td>
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 </tbody>
</table>

detection of microregional hypoxia in mouse cerebral cortex by two-photon imaging of endogenous nadh fluorescence

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. Here we present a visualized experimental and methodological protocol to directly visualize microregional tissue hypoxia and to infer perivascular oxygen gradients in the mouse cortex. It is based on the non-linear relationship between nicotinamide adenine dinucleotide (NADH) endogenous fluorescence intensity and oxygen partial pressure in the tissue, where observed tissue NADH fluorescence abruptly increases at tissue oxygen levels below 10 mmHg1. We use two-photon excitation at 740 nm which allows for concurrent excitation of intrinsic NADH tissue fluorescence and blood plasma contrasted with Texas-Red dextran. The advantages of this method over existing approaches include the following: it takes advantage of an intrinsic tissue signal and can be performed using standard two-photon in vivo imaging equipment; it permits continuous monitoring in the whole field of view with a depth resolution of ~50 &mu;m. We demonstrate that brain tissue areas furthest from cerebral blood vessels correspond to vulnerable watershed areas which are the first to become functionally hypoxic following a decline in vascular oxygen supply. This method allows one to image microregional cortical oxygenation and is therefore useful for examining the role of inadequate or restricted tissue oxygen supply in neurovascular diseases and stroke.

Watch this Scientific Journal Video about Detection of Microregional Hypoxia in Mouse Cerebral Cortex by Two-photon Imaging of Endogenous NADH Fluorescence at JoVE.com

Detection of Microregional Hypoxia in Mouse Cerebral Cortex by Two-photon Imaging of Endogenous NADH Fluorescence

Here we describe a method to directly visualize microregional tissue hypoxia in the mouse cortex in vivo. It is based on concurrent two-photon imaging of nicotinamide adenine dinucleotide (NADH) and the cortical microcirculation. This method is useful for high resolution analysis of tissue oxygen supply.

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. ...

detection-microregional-hypoxia-mouse-cerebral-cortex-two-photon

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JoVE Journal

Neuroscience

16.0K Views.  University of Rochester Medical Center. Here we describe a method to directly visualize microregional tissue hypoxia in the mouse cortex in vivo. It is based on concurrent two-photon imaging of nicotinamide adenine dinucleotide (NADH) and the cortical microcirculation. This method is useful for high resolution analysis of tissue oxygen supply.

Video: Detection of Microregional Hypoxia in Mouse Cerebral Cortex by Two-photon Imaging of Endogenous NADH Fluorescence

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has significantly broadened our knowledge of physiological and pathological processes in numerous tissues in vivo 4-7. In studies of the central nervous system (CNS), there has been a broad application of in vivo imaging in the brain, which has produced a plethora of novel and often unexpected findings about the behavior of cells such as neurons, astrocytes, microglia, under physiological or pathological conditions 8-17. However, mostly technical complications have limited the implementation of in vivo imaging in studies of the living mouse spinal cord. In particular, the anatomical proximity of the spinal cord to the lungs and heart generates significant movement artifact that makes imaging the living spinal cord a challenging task.
We developed a novel method that overcomes the inherent limitations of spinal cord imaging by stabilizing the spinal column, reducing respiratory-induced movements and thereby facilitating the use of two-photon microscopy to image the mouse spinal cord in vivo. This is achieved by combining a customized spinal stabilization device with a method of deep anesthesia, resulting in a significant reduction of respiratory-induced movements. This video protocol shows how to expose a small area of the living spinal cord that can be maintained under stable physiological conditions over extended periods of time by keeping tissue injury and bleeding to a minimum. Representative raw images acquired in vivo detail in high resolution the close relationship between microglia and the vasculature. A timelapse sequence shows the dynamic behavior of microglial processes in the living mouse spinal cord. Moreover, a continuous scan of the same z-frame demonstrates the outstanding stability that this method can achieve to generate stacks of images and/or timelapse movies that do not require image alignment post-acquisition. Finally, we show how this method can be used to revisit and reimage the same area of the spinal cord at later timepoints, allowing for longitudinal studies of ongoing physiological or pathological processes in vivo.

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

A minimally invasive protocol to stabilize the mouse spinal column and perform repetitive in vivo spinal cord imaging using two-photon microscopy is described. This method combines a spinal stabilization device and an anesthetic regimen to minimize respiratory-induced movements and produce raw imaging data that require no alignment or other post-processing.

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

Two-photon Imaging of Cellular Dynamics in the Mouse Spinal Cord

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for live-cell imaging in the murine spinal cord. This technique is uniquely suited to analyze neural precursor cell (NPC) dynamics following transplantation into spinal cords undergoing neuroinflammatory demyelinating disorders. NPCs migrate to sites of axonal damage, proliferate, differentiate into oligodendrocytes, and participate in direct remyelination. NPCs are thereby a promising therapeutic treatment to ameliorate chronic demyelinating diseases. Because transplanted NPCs migrate to the damaged areas on the ventral side of the spinal cord, traditional intravital 2P imaging is impossible, and only information on static interactions was previously available using histochemical staining approaches. Although this method was generated to image transplanted NPCs in the ventral spinal cord, it can be applied to numerous studies of transplanted and endogenous cells throughout the entire spinal cord. In this article, we demonstrate the preparation and imaging of a spinal cord with enhanced yellow fluorescent protein-expressing axons and enhanced green fluorescent protein-expressing transplanted NPCs.

A new ex vivo preparation for imaging the mouse spinal cord. This protocol allows for two-photon imaging of live cellular interactions throughout the spinal cord.

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon ...

Quantification of Cerebral Vascular Architecture using Two-photon Microscopy in a Mouse Model of HIV-induced Neuroinflammation

Human Immunodeficiency Virus 1 (HIV-1) infection frequently results in HIV-1 Associated Neurocognitive Disorders (HAND), and is characterized by a chronic neuroinflammatory state within the central nervous system (CNS), thought to be driven principally by virally-mediated activation of microglia and brain resident macrophages. HIV-1 infection is also accompanied by changes in cerebrovascular blood flow (CBF), raising the possibility that HIV-associated chronic neuroinflammation may lead to changes in CBF and/or in cerebral vascular architecture. To address this question, we have used a mouse model for HIV-induced neuroinflammation, and we have tested whether long-term exposure to this inflammatory environment may damage brain vasculature and result in rarefaction of capillary networks. In this paper we describe a method to quantify changes in cortical capillary density in a mouse model of neuroinflammatory disease (HIV-1 Tat transgenic mice). This generalizable approach employs in vivo two-photon imaging of cortical capillaries through a thin-skull cortical window, as well as ex vivo two-photon imaging of cortical capillaries in mouse brain sections. These procedures produce images and z-stack files of capillary networks, respectively, which can be then subjected to quantitative analysis in order to assess changes in cerebral vascular architecture.

This paper describes a method by which the vascular architecture in the brain can be quantified using in vivo and ex vivo two-photon microscopy.

Human Immunodeficiency Virus 1 (HIV-1) infection frequently results in HIV-1 Associated Neurocognitive Disorders (HAND), and is characterized by a chronic ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent and cause visual impairment and blindness. Understanding how such diseases progress is critical to formulating new treatments. In vivo&#160;microscopy in animal models of disease is a powerful tool for understanding neurodegeneration and has led to important progress towards treatments of conditions ranging from Alzheimer&#39;s disease to stroke. Given that the retina is the only central nervous system structure inherently accessible by optical approaches, it naturally lends itself towards in vivo imaging. However, the native optics of the lens and cornea present some challenges for effective imaging access.
This protocol outlines methods for in vivo two-photon imaging of cellular cohorts and structures in the mouse retina at cellular resolution, applicable for both acute- and chronic-duration imaging experiments. It presents examples of retinal ganglion cell (RGC), amacrine cell, microglial, and vascular imaging using a suite of labeling techniques including adeno-associated virus (AAV) vectors, transgenic mice, and inorganic dyes. Importantly, these techniques extend to all cell types of the retina, and suggested methods for accessing other cellular populations of interest are described. Also detailed are example strategies for manual image postprocessing for display and quantification. These techniques are directly applicable to studies of retinal function in health and disease.

In vivo imaging is a powerful tool for the study of biology in health and disease. This protocol describes transpupillary imaging of the mouse retina with a standard two-photon microscope. It also demonstrates different in vivoimaging methods to fluorescently label multiple cellular cohorts of the retina.

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent ...

In Vivo Chronic Two-Photon Imaging of Microglia in the Mouse Hippocampus

Microglia, the only immune cells resident in the brain, actively participate in neural circuit maintenance by modifying synapses and neuronal excitability. Recent studies have revealed the differential gene expression and functional heterogeneity of microglia in different brain regions. The unique functions of the hippocampal neural network in learning and memory may be associated with the active roles of microglia in synapse remodeling. However, inflammatory responses induced by surgical procedures have been problematic in the two-photon microscopic analysis of hippocampal microglia. Here, a method is presented that enables the chronic observation of microglia in all layers of the hippocampal CA1 through an imaging window. This method allows the analysis of morphological changes in microglial processes for more than 1 month. Long-term and high-resolution imaging of the resting microglia requires minimally invasive surgical procedures, appropriate objective lens selection, and optimized imaging techniques. The transient inflammatory response of hippocampal microglia may prevent imaging immediately after surgery, but the microglia restore their quiescent morphology within a few weeks. Furthermore, imaging neurons simultaneously with microglia allows us to analyze the interactions of multiple cell types in the hippocampus. This technique may provide essential information about microglial function in the hippocampus.

In Vivo Chronic Two-Photon Imaging of Microglia in the Mouse Hippocampus

This paper describes a method for chronic in vivo observation of the resting microglia in the mouse hippocampal CA1 using precisely controlled surgery and two-photon microscopy.