This work was supported by grants from the National Institutes of Health (1R21NS094061-01A1) (NIH 1R01EB012874-01) (S10RR031637), and the National Science Foundation (cooperative agreement No. DMR-1157490) through the National High Magnetic Field Laboratory (NHMFL) Advanced Magnetic Resonance Imaging and Spectroscopy (AMRIS) facility at UF and the State of Florida.

All animal experiments described follow the guidelines set forth in the National Academies of Sciences&#39; Guide for the Care and Use of Laboratory Animals and were reviewed and approved by the University of Florida&#39;s Institutional Animal Care and Use Committee (IACUC). Follow all applicable rules and regulations when engaging in animal subject research.
1. Preparation of Perfusate for the Maintenance of Central Nervous System Tissues
<ol>
	<li>Make fresh artificial cerebrospinal fluid (aCSF).
	<ol>
		<li>To generate 2 L of bicarbonate-buffered aCSF perfusate, measure out 1,500 mL of water purified by double distillation or reverse osmosis into a 4 L flask. Place a magnetic stir bar in the flask and agitate the liquid using a stir plate.</li>
		<li>Into the purified water, dissolve the following quantities of salts: 14.03 g (120 mM) sodium chloride, 4.37 g (26 mM) sodium bicarbonate, 0.41 g (1.5 mM) monobasic potassium phosphate, 0.69 g (1.4 mM) magnesium sulfate heptahydrate, 0.59 g (2 mM) calcium chloride dihydrate, 0.45 g (3 mM) potassium chloride, and 3.6 g (10 mM) glucose. 
		NOTE: The chemical contents of the perfusate will differ depending on the specific metabolic requirements of the tissue and the desired conditions of the experiment.</li>
		<li>Mix this solution thoroughly until all salts have dissolved. Adjust volume to 2 L using additional purified water.</li>
		<li>Test the osmolality of the aCSF using a freezing point depression osmometer. Adjust to 300 mOsm/kg using sorbitol. Addition of 324 mg sorbitol to 2 L of 299 mOsm/kg aCSF perfusate will increase osmolality to 300 mOsm/kg (approximately 1 mOsm per 324mg). 
		NOTE: The aCSF can be stored at 4 &#176;C in an air-tight, sealed bottle for a period not to exceed 24 h.</li>
	</ol>
	</li>
</ol>
2. Set Up the Perfusion System
<ol>
	<li>Prepare the aCSF perfusate.
	<ol>
		<li>Equilibrate aCSF to ambient temperature (~23 &#176;C) while gassing with 95% carbogen (95% O2, 5% CO2; 1/16 L/min) via direct bubbling for no less than 1 h. 
		NOTE: Different tissue types or desired experimental conditions may require adjustment of the perfusate temperature or gas contents. The desired conditions for dissolved gas content in the perfusate can be controlled precisely by varying the percent concentrations of individual components of the supply gas (Figure 1).</li>
		<li>Once the aCSF reaches the required temperature and carbogen saturation, confirm that the pH is in the physiological range (7.3 - 7.4). Continue to bubble gas directly into the perfusate reservoir (1/16 L/min) throughout the course of experiment in order to maintain proper dissolved oxygen content and pH conditions conducive to healthy tissue metabolism.</li>
	</ol>
	</li>
	<li>Prime the perfusion lines.
	<ol>
		<li>Submerge the inlet tube connected to the peristaltic micro-pump into the prepared aCSF (300 mOsm, pH 7.3 - 7.4, continuously gassed).</li>
		<li>Pass the oxygenator device (Figure 2) and perfusion lines through both the magnet bore and gradient coil stack (top to bottom) and set the coils aside until probe assembly. Hang the oxygenator above a beaker to catch any aCSF effluent. 
		NOTE: Depending on the MRI system design, perfusion lines may need to be passed through the magnet bore and gradient coils or probe body base prior to priming the system with aCSF. Confirm that perfusion line placement will not interfere with the assembly of the probe body or insertion of the probe into the bore before beginning the priming procedure.</li>
		<li>Confirm the intended flow rate (2 mL/min) is selected and begin filling the perfusion lines by switching on the pump.</li>
		<li>Invert the empty, in-line bubble trap so that aCSF will displace the air contained inside.</li>
		<li>Connect the oxygenator&#39;s gas port to a second source of carbogen (either a manifold or a secondary carbogen cylinder) and set a flow rate of 1/16L/min.</li>
		<li>Confirm that carbogen is flowing over the oxygenator&#39;s gas-exchange membrane by dipping the exhaust port on top of the oxygenator into water.</li>
		<li>Once aCSF is detected dripping from the perfusion chamber, purge any visible gas bubbles from the inflow lines by manual agitation.</li>
		<li>Confirm the oxygenator is operating correctly by submerging the oxygen electrode in the aCSF flowing from the perfusion chamber. 
		NOTE: The reading on the oxygen meter should approximate the percentage of oxygen contained within the supplied gas.</li>
	</ol>
	</li>
</ol>
3. Tissue Preparation
<ol>
	<li>Euthanasia
	<ol>
		<li>Place a rat (125 - 150 g) into the induction chamber of an anesthesia machine and expose to 4% isoflurane in an oxygen carrier gas at a rate of 1 L/min for 4 min.</li>
		<li>Remove the unconscious rat from the anesthesia chamber.</li>
		<li>Perform a righting reflex test by placing the rat in the supine position. Check for any movements&#8212;head turning, leg lifting, spine flexing, etc.&#8212;indicative that the animal is turning onto its stomach.</li>
		<li>Perform an eye-blink reflexive test by touching the open eye of the animal using a cotton swab. Check for any response&#8212;lid closure, muscle twitching, etc.&#8212;to this sensory input.</li>
		<li>Lastly, perform a limb-withdraw reflex test by pinching the skin between the toes of the rat&#39;s outstretched back leg using tweezers or hemostats. Check for flexion of the leg. 
		NOTE: In the case of a positive reflexive test result, do not proceed with euthanasia.</li>
		<li>In the event that any of the three reflex tests elicits a response, return the rat immediately to the anesthesia chamber and allow for an additional 2 min of exposure to 4% isoflurane.</li>
		<li>Perform all three reflexive tests over in their entirety. Repeat the 2 min anesthesia exposure as necessary and only proceed once a complete lack of response to all three reflexive tests has been observed.</li>
		<li>Euthanize the rat via guillotine decapitation.</li>
	</ol>
	</li>
	<li>Brain resection
	<ol>
		<li>Remove rat brain by gross dissection. Start with the head in the prone position. Using scissors, cut rostrally through the skin from the back of the neck to the nose and expose the skull.</li>
		<li>Remove soft tissues from the surface of the skull with a blunt probe.</li>
		<li>Working from the caudal end, remove the occipital, parietal, and frontal bones of the cranium using rongeurs.</li>
		<li>Retract the dura by cutting along the longitudinal fissure with micro scissors and peeling back the layer from both hemispheres.</li>
		<li>Remove the brain from the skull by turning the head to the supine position and severing the cranial nerves on the ventral side.</li>
	</ol>
	</li>
	<li>Slice isolation
	<ol>
		<li>Return the brain to the prone position and isolate the central portion containing the hippocampus by removing all tissues caudal to the transverse fissure and rostral to the fimbria. Make two cuts along the transverse plane (i.e. coronal slices) with a straight-edged razor.</li>
		<li>Affix the brain&#39;s caudal-most plane to the center of a vibratome cutting bath using cyanoacrylate glue.</li>
		<li>Add ice-cold, carbogen bubbled aCSF to the vibratome cutting bath and place the nylon retention hardware inside.</li>
		<li>Cut 300 &#956;m thick slices to obtain 3 to 4 usable slice preparations per hemisphere (6 to 8 total).</li>
		<li>Isolate a hippocampus or cortical section from one hemisphere and trim the slice so that it fits within the microcoil&#39;s 5 mm diameter tissue well.</li>
	</ol>
	</li>
</ol>
4. Sample Positioning and Perfusion System Assembly
<ol>
	<li>Coil preparation
	<ol>
		<li>Fill the microcoil&#39;s tissue chamber with oxygenated aCSF from the vibratome&#39;s cutting bath using a transfer pipette.</li>
		<li>Take the trimmed brain sample and position the region of interest (eg. pyramidal cell layer) over the microcoil using a dissecting scope.</li>
		<li>Insert the tissue retention device (nylon mesh net affixed to nylon washer) to retain sample position throughout the imaging experiment. 
		NOTE: Proceed quickly through the sample positioning procedure to minimize exposure to intense light. Provide additional oxygenated aCSF as needed using a transfer pipette.</li>
	</ol>
	</li>
	<li>Assembling the in-bore oxygenator, microcoil and probe
	<ol>
		<li>Secure the modified microcoil assembly (Figure 3) in a table clamp and affix the in-bore oxygenator device by inserting the acetal support peg into the hole on top of the coil.</li>
		<li>Seal the perfusion system by placing the perfusion chamber over the microcoil&#39;s tissue well and cinching the two together using a miniature cable tie.</li>
		<li>Trim the excess length from the cable tie using wire cutters. 
		NOTE: Upon successful sealing, perfusate should be observed exiting through the outflow lines and no leakage should be evident around the silicone seal of the perfusion chamber. Failure to confirm these conditions prior to probe assembly could result in serious damage to imaging hardware.</li>
		<li>Once aCSF can be seen dripping into the waste reservoir, take the microcoil and oxygenator and attach the assembly to the top of the imaging probe body.</li>
		<li>Slide the gradient stack over the assembly and seat the gradient on top of the probe. Photographs detailing the relative placement and proper connection of coil, oxygenator and probe hardware components are provided (Figure 4).</li>
	</ol>
	</li>
</ol>
5. Performing the MR Image Collection
<ol>
	<li>Inserting the assembled probe into the magnet bore
	<ol>
		<li>Place the probe body in close proximity to the spectrometer bore opening at the bottom of the magnet.</li>
		<li>Retract the excess length of perfusion lines through the bore opening at the top of the magnet.</li>
		<li>Once all of the available slack has been taken up from the perfusion lines, advance the probe body into the magnet bore at the base while simultaneously removing more slack from the perfusion lines from the top of the bore.</li>
		<li>Thread the two securing screws at the base of the probe into the corresponding slots of the shim stack.</li>
		<li>Before proceeding, check that perfusion lines were not pinched or kinked during probe insertion by confirming aCSF outflow into the waste reservoir. 
		NOTE: Failure to confirm perfusate outflow can result in permanent damage to perfusion lines and risks catastrophic damage to MR imaging hardware.</li>
		<li>In the event no perfusate is seen exiting the return line into the waste reservoir, remove the probe body and confirm that perfusate flow has recommenced prior to attempting reinsertion. A schematic layout of the assembled perfusion system and imaging spectrometer has been described previously8.</li>
	</ol>
	</li>
	<li>Connecting the Probe Body
	<ol>
		<li>Attach the inflow and outflow water lines from the gradient chiller.</li>
		<li>Turn on the pump for gradient chiller and confirm the water temperature setting (19 &#176;C).</li>
		<li>Attach the air hose from the air chiller unit to the probe body using a hose clamp.</li>
		<li>Turn the flow knob on the air chiller unit to the &#34;1&#34; position.</li>
		<li>Attach the thermocouple wire to the probe body.</li>
		<li>Attach the coaxial cable from the preamp to the radio frequency (RF) input/output on the probe body&#39;s proton (H) channel.</li>
		<li>Attach the power cable from the gradient amplifiers to the probe body.</li>
		<li>Inside the RF cabinet, turn on the power to the B0 compensation unit, all three gradient amplifiers (x,y,z), and the master unit.</li>
	</ol>
	</li>
	<li>Preparing the Spectrometer
	<ol>
		<li>Set the intended bore temperature using the variable temperature adjustment module at the console.</li>
		<li>Match (impedance) and tune (frequency) the RF circuit by adjusting the variable capacitors within the probe. This is accomplished by manipulating the wands at the base of the probe body.</li>
		<li>Adjust the current settings for the spectrometer&#39;s shim coils to maximize the magnetic field homogeneity at the sample.</li>
	</ol>
	</li>
	<li>Begin Image Collection
	<ol>
		<li>Collect pilot images to determine the spatial position of your sample within the magnet bore. Typical parameters for a two dimensional gradient echo are as follows (TR/TE = 100/4 ms, averages = 1, pulse angle = 30o, time = 6 sec, matrix = 64 x 64, field of view = 0.3 x 0.3 cm, resolution = 47 x 47 &#956;m).</li>
		<li>Collect diffusion-weighted pilots in order to confirm proper scan geometry and tissue position if applicable. Typical parameters for a two dimensional diffusion-weighted pilot scan are as follows (TR/TE = 2000/11.6 ms, time = 4.3 min, &#916; = 6 ms, &#948; = 1 ms, averages = 1, b = 1200 (1860 effective) s/mm2, matrix = 64 x 64, field of view = 0.2 x 0.2 cm, resolution = 31 &#956;m).</li>
		<li>Collect a diffusion-weighted time series to determine the stability characteristics of the acute slice preparation. Typical parameters for a two dimensional diffusion weighted image are as follows (TR/TE = 2000/11.6 ms, time = 1.5 h, &#916; = 6 ms, &#948; = 1 ms, averages = 42, b = 1200 s/mm2, matrix = 64 x 64, field of view = 0.2 x 0.2 cm, resolution = 31 &#956;m). Note: Characterizing stability in a given system will vary from the described protocol depending on factors such as the MR contrast employed (e.g. T1,T2, diffusion, susceptibility), the physical perturbation studied, and the MR signal change per unit time resulting from said perturbation.</li>
	</ol>
	</li>
</ol>

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	<li>Aguayo, J. B., Blackband, S. J., Schoeniger, J., Mattingly, M. A., Hintermann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nuclear+magnetic+resonance+imaging+of+a+single+cell.">Nuclear magnetic resonance imaging of a single cell.</a> Nature. 322, 190-191 (1986).</li><li>Schoeniger, J. S., Aiken, N., Hsu, E., Blackband, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relaxation-time+and+diffusion+NMR+microscopy+of+single+neurons.">Relaxation-time and diffusion NMR microscopy of single neurons.</a> J. Magn. Reson. B. 103, 261-273 (1994).</li><li>Flint, J. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Magnetic+resonance+microscopy+of+mammalian+neurons.">Magnetic resonance microscopy of mammalian neurons.</a> Neuroimage. 46, 1037-1040 (2009).</li><li>Flint, J. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Magnetic+resonance+microscopy+of+human+and+porcine+neurons+and+cellular+processes.">Magnetic resonance microscopy of human and porcine neurons and cellular processes.</a> Neuroimage. 60, 1404-1411 (2012).</li><li>Kamman, R. L., Go, K. G., Stomp, G. P., Hulstaert, C. E., Berendsen, H. 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=Changes+of+Relaxation+times+T1+and+T2+in+rat+tissues+after+biopsy+and+fixation.">Changes of Relaxation times T1 and T2 in rat tissues after biopsy and fixation.</a> Magn. Reson. Imag. 3, 245-250 (1985).</li><li>Shepherd, T. M., Thelwall, P. E., Stanisz, G. J., Blackband, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aldehyde+fixative+solutions+alter+the+water+relaxation+and+diffusion+properties+of+nervous+tissue.">Aldehyde fixative solutions alter the water relaxation and diffusion properties of nervous tissue.</a> Magn. Reson. Med. 62, 26-34 (2009).</li><li>Massin, C., Boero, G., Vincent, F., Abenhaim, J., Besse, P. -. A., Popovic, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-Q+factor+RF+planar+microcoils+for+micro-scale+NMR+spectroscopy.">High-Q factor RF planar microcoils for micro-scale NMR spectroscopy.</a> Sensor. Actuat. A-Phys. 97, 280-288 (2002).</li><li>Flint, J., Menon, K., Hansen, B., Forder, J., Blackband, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+microperfusion+and+in-bore+oxygenator+system+designed+for+magnetic+resonance+microscopy+studies+on+living+tissue+explants.">A microperfusion and in-bore oxygenator system designed for magnetic resonance microscopy studies on living tissue explants.</a> Sci. Rep. 5, 18095 (2015).</li><li>Khong, Y. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+intra-tissue+perfusion+system+for+culturing+thick+liver+tissue.">Novel intra-tissue perfusion system for culturing thick liver tissue.</a> Tissue Eng. 13 (9), 2345-2356 (2007).</li><li>Schumacher, K., Khong, Y. -. M., Chang, S., Ni, J., Sun, W., Yu, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perfusion+culture+improves+the+maintenance+of+cultured+liver+tissue+slices.">Perfusion culture improves the maintenance of cultured liver tissue slices.</a> Tissue Eng. 13 (1), 197-205 (2007).</li><li>Shepherd, T. M., Blackband, S. J., Wirth, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+diffusion+MRI+measurements+from+multiple+perfused+rat+hippocampal+slices.">Simultaneous diffusion MRI measurements from multiple perfused rat hippocampal slices.</a> Magn. Reson. Med. 48, 565-569 (2002).</li><li>Henry, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experiments+on+the+quantity+of+gases+absorbed+by+water,+at+different+temperatures,+and+under+different+pressures.">Experiments on the quantity of gases absorbed by water, at different temperatures, and under different pressures.</a> Phil. Trans. R. Soc. Lond. 93, (1803).</li><li>Bui, J. D., Buckley, D. L., Phillips, M. I., Blackband, S. J., Bl&#252;mler, P., Bl&#252;mich, B., Botto, R., Fukushima, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studies+of+perfused+brain+slices+with+MR+microscopy.">Studies of perfused brain slices with MR microscopy.</a> Spatially Resolved Magnetic Resonance. , 337-343 (1998).</li><li>Moseley, M. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+detection+of+regional+cerebral+ischemia+in+cats:+Comparison+of+diffusion-+and+T2-weighted+MRI+and+spectroscopy.">Early detection of regional cerebral ischemia in cats: Comparison of diffusion- and T2-weighted MRI and spectroscopy.</a> Magn. Reson. Med. 14 (2), 330-346 (1990).</li><li>Moseley, M. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diffusion-weighted+MR+imaging+of+acute+stroke:+Correlation+with+T2-weighted+and+magnetic+susceptibility-enhanced+MR+imaging+in+cats.">Diffusion-weighted MR imaging of acute stroke: Correlation with T2-weighted and magnetic susceptibility-enhanced MR imaging in cats.</a> Am. J. Neuroradiol. 11, 423-429 (1990).</li></ol>

The authors have no conflicts of interest to disclose.

The current protocol describes procedures necessary for standard metabolic maintenance of acute brain slice preparations undergoing magnetic resonance microscopy. This procedure is the only method currently available that allows visualization of living mammalian tissues with MR at resolutions capable of resolving cells. While the perfusate conditions described are tailored specifically to central nervous system tissues, the protocol is widely adaptable to any manner of living tissue preparation through adjustments of perfusate and gas constituents as well as perfusion flow rate and temperature.
The most common problems likely to be encountered during the described procedures include those related to failures in metabolite supply. Precipitation of calcium salts can occur inside the aCSF during gaseous insufficiency as a result of failures in the bicarbonate buffering system. Such precipitates can clog the perfusion lines and result in severe hardware damage. If salt precipitates are observed in the perfusate following probe assembly, cease perfusion flow immediately by turning off the peristaltic pump. Confirm presence of sufficient sodium bicarbonate levels (4.37 g / 2 L) in perfusate, CO2 levels (5.0%) in supply gas, and carbogen gas flow (1/16 L/min) into both reservoir and oxygenator. Finally, confirm pH levels are stabilized in the physiological range (7.3 - 7.4). In the event that oxygen gas and pH levels are still not regulated appropriately, the gas-exchange membrane should be replaced.
If slices do not exhibit signal stability over the intended experimental time-course, confirm that the correct chemical constituents are present in the aCSF mixture and that the correct osmolality (300 mOsm) and pH (7.3 - 7.4) are maintained. Also, ensure carbogen gas is being supplied to the perfusate reservoir and oxygenator at 1/16 L/min. If these steps do not correct perfusate conditions, replacement of the gas-exchange membrane is advised. If tissue stability is not achieved after troubleshooting the perfusate conditions, consider refinement of the surgical protocol with a focus on minimizing the time interval between tissue harvest and perfusion application.

As magnetic resonance imaging (MRI) systems have steadily progressed to ever-higher field strengths, more details about the composition and status of living tissues have become discernable. Despite such hardware advances, MR imaging at resolutions sufficient to visualize the cellular structures of tissues is still not available in the clinic. As a result, cellular-level characteristics of tissues must be inferred when considering the content of clinical scans. Such inference requires knowledge of equivalent processes gleaned from data taken in model systems which can be observed directly. Traditionally, these models included cells from aquatic organisms such as the Xenopus laevis oocyte and Aplysia californica L7 neuron1,2. These were among the first animal cells available for observation with MR methods due to their atypically large size: approximately 1000 &#956;m and 300 &#956;m diameter, respectively. More recently, advances in hardware design have allowed for one of the largest examples of mammalian cells&#8212;the &#945;-motor neuron&#8212;to be imaged using MR microscopy techniques on fixed tissue3,4. While these studies demonstrated direct visualization of mammalian cellular material using MR, the fixed samples employed differ significantly in their MR properties from live tissue and thus cannot serve as an equivalent representative model5,6. More importantly, observing MR contrast changes that occur in concert with complex biological processes requires living samples that can be perturbed and measured over the course of the imaging experiment.
To facilitate MR microscopy studies on living tissues, a protocol is presented which includes commercial microimaging hardware7 interfaced to a purpose-built, MR compatible, in-bore oxygenator and perfusion device previously described8. Unique advantages of this design include cellular-level resolution capabilities in mammalian tissues and precision control over dissolved gas content and pH at the site of tissue perfusion. Also, unlike the majority of explant MR studies which interrupt perfusion during image acquisition to avoid flow artefacts, this design supports the use of continuous perfusion during data collection which has been shown to improve the physiological condition of isolated tissues9,10. Lastly, its closed recording chamber and slice-retention hardware aid in reducing the likelihood of motion artifacts which could otherwise occur during protracted image collection.
While the current protocol describes procedures appropriate for use with acute hippocampal and cortical slices, precise control over perfusate metabolites enables this system to accommodate a wide array of diverse tissue types and experimental conditions. Limitations of this design include a reduction in sample throughput as compared to a multi-slice perfusion chamber11; however, this limitation may be overcome in the future using multi-coil arrays.
Also, while the described system can be employed in both horizontal or vertical configurations, the current protocol features its use in a vertically oriented, 600 MHz spectrometer. Any system capable of MR microimaging studies&#8212;typically narrow-bore (&#8804;6 cm), high field (&#8805;500 MHz) spectrometers&#8212;will accommodate the oxygenator and perfusion equipment described. However, changes to the imaging coil, gradient, probe system, or other essential imaging hardware employed may necessitate alterations to the perfusion equipment and MR scan parameters.

<table>
	<tbody>
		<tr>
			<td>Perfusate Preparation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Osmette A</td>
			<td>Precision Systems Inc.</td>
			<td>5002</td>
			<td>freezing point depression osmometer</td>
		</tr>
		<tr>
			<td>Stir Plate Type 1000</td>
			<td>Barnstead/Thermodyne</td>
			<td>SPA1025B</td>
			<td>magnetic stir plate with heating element</td>
		</tr>
		<tr>
			<td>Accumet Basic pH Meter</td>
			<td>Fisher Scientific</td>
			<td>AB15</td>
			<td>pH Meter</td>
		</tr>
		<tr>
			<td>pH Probe</td>
			<td>Fisher Scientific</td>
			<td>13-620-AP61</td>
			<td>probe for pH measurement</td>
		</tr>
		<tr>
			<td>Oxygen Meter</td>
			<td>Microelectrodes Inc.</td>
			<td>OM-4</td>
			<td>meter for sampling the oxygen content of gasses or the disolved oxygen content of liquid perfusates</td>
		</tr>
		<tr>
			<td>Oxygen Electrode</td>
			<td>Microelectrodes Inc.</td>
			<td>MI-730</td>
			<td>microprobe for the oxygen meter</td>
		</tr>
		<tr>
			<td>Scale</td>
			<td>Denver Instrument Co.</td>
			<td>A-160</td>
			<td>microscale for weighing chemical components</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Slice Preparation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Lancer Vibratome</td>
			<td>Ted Pella Inc.</td>
			<td>Series 1000</td>
			<td>vibratory tissue slicer</td>
		</tr>
		<tr>
			<td>Disecting Microscope</td>
			<td>Carl Zeiss Inc.</td>
			<td>OPMI 1-FC</td>
			<td>tabletop, binocular disecting microscope</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Perfusion System</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Masterflex L/S</td>
			<td>Cole-Parmer</td>
			<td>7523-50</td>
			<td>peristaltic micro perfusion pump</td>
		</tr>
		<tr>
			<td>Oxygen Regulators x 2</td>
			<td>Victor Medical</td>
			<td>VMG-05LY</td>
			<td>device for regulating gas flow</td>
		</tr>
		<tr>
			<td>e-sized carbogen cylinders x 2</td>
			<td>Airgas</td>
			<td></td>
			<td>gas tanks containing carbogen gas</td>
		</tr>
		<tr>
			<td>in-bore oxygenator</td>
			<td>developed in house</td>
			<td></td>
			<td>device responsible for pH and oxygen regulation in the perfusate</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>MR Imaging Hardware</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Micro Surface Coil (200mm dia., modified)</td>
			<td>Bruker Biospin</td>
			<td>B6371/0001</td>
			<td>four-turn micro (200mm dia) surface-style radiofrequency coil</td>
		</tr>
		<tr>
			<td>Micro 5 probe body</td>
			<td>Bruker Biospin</td>
			<td>Z3395</td>
			<td>microimaging probe used in the 600 MHz spectrometer</td>
		</tr>
		<tr>
			<td>Micro 5 gradient coils</td>
			<td>Bruker Biospin</td>
			<td>M81111</td>
			<td>gradient coil stack used with micro 5 probe body</td>
		</tr>
		<tr>
			<td>600 MHz Spectrometer</td>
			<td>Oxford Instruments</td>
			<td></td>
			<td>superconducting magnet (14.1T) used for MR image generation</td>
		</tr>
		<tr>
			<td>Imaging Console</td>
			<td>Bruker Biospin</td>
			<td>Avance III</td>
			<td>support and control hardware including gradient amplifiers, preamps, &#38; workstation used for MR image generation</td>
		</tr>
		<tr>
			<td>Air Blower</td>
			<td>Bruker Biospin</td>
			<td>BCU-II, -80/60</td>
			<td>Air chiller unit used in conjunction with the probe&#39;s heating coil to regulate temperature inside the magnet bore</td>
		</tr>
		<tr>
			<td>Gradient Chiller</td>
			<td>Thermo Scientific</td>
			<td>Neslab Merlin M33</td>
			<td>Water chiller used to disipate heat generated by the gradient coils</td>
		</tr>
	</tbody>
</table>

metabolic support of excised, living brain tissues during magnetic resonance microscopy acquisition

This protocol describes the procedures necessary to support normal metabolic functions of acute brain slice preparations during the collection of magnetic resonance (MR) microscopy data. While it is possible to perform MR collections on living, excised mammalian tissue, such experiments have traditionally been constrained by resolution limits and are thus incapable of visualizing tissue microstructure. Conversely, MR protocols that did achieve microscopic image resolution required the use of fixed samples to accommodate the need for static, unchanging conditions over lengthy scan times. The current protocol describes the first available MR technique that enables imaging of living, mammalian tissue samples at microscopic resolutions. Such data is of great importance to the understanding of how pathology-based contrast changes occurring at the microscopic level influence the content of macroscopic MR scans such as those used in the clinic. Once such an understanding is realized, diagnostic methods with greater sensitivity and accuracy can be developed, which will translate directly to earlier disease treatment, more accurate therapy monitoring and improved patient outcomes.
While the described methodology focuses on brain slice preparations, the protocol is adaptable to any excised tissue slice given that changes are made to the gas and perfusate preparations to accommodate the tissue&#39;s specific metabolic needs. Successful execution of the protocol should result in living, acute slice preparations that exhibit MR diffusion signal stability for periods up to 15.5 h. The primary advantages of the current system over other MR compatible perfusion apparatuses are its compatibility with the MR microscopy hardware required to attain higher resolution images and ability to provide constant, uninterrupted flow with carefully regulated perfusate conditions. Reduced sample throughput is a consideration with this design as only one tissue slice may be imaged at a time.

Perfusate Preparation
Upon successful employment of the in-bore oxygenation device, gases present in the supplied carbogen will reach 100% saturation conditions within the aCSF perfusate. This can be demonstrated by varying the oxygen concentration of the supplied gas and measuring the change in dissolved oxygen content in the aCSF perfusate within the perfusion chamber using an oxygen meter (Figure 1)8. According to Henry&#39;s law, the amount of dissolved gas that is in equilibrium with a liquid sample is directly proportional to the partial pressure of that gas provided that the temperature remains constant12. Using this knowledge and precision gas standards, it is possible to quantify the amount of dissolved oxygen contained within an aCSF sample as described. This is achieved by calibrating the oxygen meter using saturated solutions (directly bubbled for 1 h&#160;or more) of aCSF being exposed to gasses of known composition: one gas with high oxygen concentration such as carbogen (95% O2) and another with low oxygen concentration such as nitrogen (0% O2). Afterward, measurements can be taken by submerging the tip of the oxygen electrode into a sample. Confirmation that the in-bore oxygenator is functioning properly can be achieved by measuring the effluent from the perfusion well. The percent dissolved oxygen as measured by the oxygen meter should match the percent concentration of oxygen delivered in the supply gas. If the measured values are lower than those in the supply gas, this would suggest a hardware failure that could lead to metabolic insufficiency in the tissue slice.
Sample Appearance and Behavior
Acute slice preparations that receive perfusion sufficient enough to supply necessary metabolites and carry away metabolic wastes soon reach a state of relative stability. From that point, acute slices can be subjected to external perturbation and their responses to these changes can be measured for scientific study. For MR experiments, tracking the signal of interest over time is a commonly used practice to demonstrate the relative stability of acute slice preparations13. The diffusion-weighted signal is especially sensitive to changes in a tissue&#39;s water mobility, content, and distribution, as can be appreciated by the use of this contrast mechanism to detect infarcts in ischemic stroke14,15. Plotting the normalized diffusion signal over time in acute cortical slices maintained under a variety of perfusion conditions demonstrates relatively stability (2 &#177; 3% over 15.5 h) after tissue isolation is achieved (Figure 5). Diffusion signal stability was maintained regardless of perfusion conditions (intermittent or continuous) or MRI scan length (short [4 min] or long [1.5 h])8. If slices do not exhibit signal stability over time, such as the sharp diffusion signal increase observed in living cortex that did not receive perfusion, this is suggestive of suboptimal experimental conditions. Perturbation experiments should not be attempted prior to confirmation of stable signal conditions in slice preparations.
In addition to signal stability, correct sample positioning must be confirmed at the time of image collection. Even though sample position is controlled during tissue placement at the dissecting microscope, shifts in sample position may occur during the assembly of the perfusion apparatus, or due to rough handling of the coil or probe prior to insertion into the magnet. Confirmation of proper hippocampal placement can be achieved by collecting short (2 min), pilot scans with diffusion contrast (Figure 6). Because the pyramidal cell layer is more sensitive to diffusion weighting than the adjacent hippocampal laminae, this structure will appear as a darker band in diffusion-weighted images. Setups that do not display this characteristic feature contain off-center samples and will likely need to be repeated.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56282/56282fig1.jpg" /> 
Figure 1: Dissolved oxygen content of aCSF perfusate as a function of percent O2 content in supplied gas. Carbogen mixtures containing variable concentrations of oxygen (95%, 60%, and 19%) are employed as a supply gas. Percent dissolved oxygen readings are then taken from the perfusion well and compared to two known controls: a perfusate reservoir directly bubbled with carbogen (95% O2), and a perfusate reservoir exposed to atmospheric conditions (23% O2). In each case, the percent oxygen saturation at the site of tissue perfusion approaches 100% of the O2 concentration within the carbogen mixture used. Error bars are equal to the standard deviation of the sample means. Figure reproduced with permission from the original article8. <a href="https://cloudflare.jove.com/files/ftp_upload/56282/56282fig1large.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/56282/56282fig2.jpg" /> 
Figure 2: Schematic drawing of the in-bore oxygenator and perfusion chamber. This diagram shows detailed design elements responsible for the function of these critical devices. Fresh perfusate that has been pumped through a bubble trap enters the oxygenator through the top of a 10 mm NMR tube. In doing so, it transitions into a highly gas permeable silicone tubing (blue segment) that is coiled around an open-ended, 5 mm NMR tube nested inside. Carbogen gas supplied through the top of the 5mm tube enters the chamber through the open-ended bottom and passes over the coiled silicone tubing before exiting the oxygenator through a vent hole in the 10 mm tube cap. During this exposure, the perfusate flowing through the silicone tube becomes saturated with the chemical components of the supplied gas mixture. Upon exiting the oxygenator, perfusate passes directly into the perfusion chamber before entering the return line leading to a waste collection reservoir. Other components critical to this design include the acetal support peg which allows the oxygenator to stand vertically atop the modified RF microcoil, a silicone washer (red ring) which forms the liquid-tight seal between the oxygenator&#39;s perfusion chamber and the microcoil&#39;s tissue well, and the cable tie notch which accommodates placement of a cable tie used to form this reversible seal. This figure has been modified and reproduced with permission from the original article8. <a href="https://cloudflare.jove.com/files/ftp_upload/56282/56282fig2large.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/56282/56282fig3.jpg" /> 
Figure 3: Modifications to the microcoil assembly which permit interface to the in-bore oxygenator. Two grooves 3.0 mm x 1.5 mm (black arrows) were cut into the side of the assembly that accommodate the width of a cable tie used to seal the perfusion chamber. A channel (15 mm x 3 mm x 4 mm) connects the groves across the back face of the coil. Two nylon spaces placed in the sides of the channel (red arrows) act as a catch for the cable tie head which eases the sealing procedure. A hole (2 mm x 14 mm) drilled in the top of the coil assembly (yellow arrow) joins to the acetal support peg to secure the oxygenator. This figure has been modified and reproduced with permission from the original article8. <a href="https://cloudflare.jove.com/files/ftp_upload/56282/56282fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56282/56282fig4.jpg" /> 
Figure 4: Photo montage detailing the relative placement and proper assembly of microcoil, oxygenator, and probe components. These images feature key hardware components of the in-bore oxygenator and microperfusion device and illustrate how the separate parts interface with one another. (A) Exploded-view photo showing the relative placement of all components prior to sealing of the tissue well or assembly of the probe body. Care has been taken to display the relative location of parts accurately; however, a section of the perfusion lines has been doubled back in this series so that all components fit within the image frame. (1 = probe head, 2 = microcoil assembly, 3 = nylon tissue retention ring, 4 = perfusion well, 5 = cable tie, 6 = in-bore oxygenator, 7 = gradient coils, 8 = bubble trap). (B) Components following coil and oxygenator assembly. In this image, the nylon retention ring has been placed within the microcoil&#39;s tissue well to secure a sample. The acetal support peg on the base of the oxygenator has been secured in the corresponding hole on top of the microcoil. The silicone gasket on the open end of the perfusion well has been placed over the tissue well, and a cable tie has been tightened around these components to seal the perfusion chamber. Lastly, the base of the microcoil has been connected to the top of the probe head. (C) Components following probe assembly. In the last panel, excess length from the cable tie has been trimmed flush with the microcoil. The gradient coil stack is then slid into position by carefully advancing the cylinder towards the probe while passing the excess perfusion lines, oxygenator, and microcoil through its hollow center. Once the gradients are connected to the probe head, they&#39;re held in place by screwing the securing collar on the probe over the threaded base of the gradients. <a href="https://cloudflare.jove.com/files/ftp_upload/56282/56282fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56282/56282fig5.jpg" /> 
Figure 5: Diffusion signal stability in superfused acute cortical slices. (A) Normalized diffusion signal values in four acute slices subjected to differing superfusion paradigms are plotted over time for a period of up to 21.5 h following euthanasia. Slices remain within &#177; 5% of their initial diffusion signal measurement for a period of 15.5 h following euthanasia regardless of whether superfusion is continuous or intermittent and independent of the MR scan length (1.5 h or 4 min). Signal recordings taken from formaldehyde-fixed cortex serve as the positive control (n = 1) for stability due to the static, unchanging nature of fixed tissue samples. Conversely, diffusion signal measured in a live slice absent of superfusion support (n = 1) serves as a control for metabolic deficiency. Experiment parameters of the different superfusion trials are as follows: continuous (superfusion always on, time per scan = 1.5 h), intermittent (superfusion on for 10 min interval between scans, time per scan = 1.5 h), Long interval, long scan (superfusion on during scan, but paused for 10 min between scans, time per scan = 1.5 h), Long interval, short scan (superfusion on for 1.5 h interval between scans, time per scan = 4 min). (B) Analyzed data showing group means of the four live-slice superfusion experiments from panel (A). The diffusion signal profile from the grouped, superfused cortical slices exhibits little variation over time (2 &#177; 3% over 15.5 h) whereas the non-perfused control (n = 1) exhibits dramatic signal instability early in the experiment (15% by 6.5 h). This figure has been modified and reproduced with permission from the original article8. <a href="https://cloudflare.jove.com/files/ftp_upload/56282/56282fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56282/56282fig6.jpg" /> 
Figure 6: Confirmation of hippocampal slice placement during pilot imaging. Prior to running an extended MR microscopy session, correct placement of the sample is critical to ensure resources such as scanner time and expensive perfusate additives are not wasted. The pyramidal cell layer in the CA1 region of the hippocampus can be visualized in faster (4.3 min), lower resolution (31 &#956;m x 31 &#956;m in-plane) pilot scans to ensure the tissue of interest is placed correctly in relation to the micro-coil. Scan parameters common to both images are as follows: TR/TE = 2000/11.6 ms, &#916; = 6 ms, &#948; = 1 ms, averages = 1. (A) b = 0 (227 effective) s/mm2. In this preliminary scan, the stratum pyramidale is only just visible as a grey, diagonal band centered in the coil&#39;s excitation profile. (B) b = 1,200 (1,860 effective) s/mm2. At higher diffusion weighting, interlamellar contrast increases as the pyramidal cell layer becomes darker than tissues in the adjacent laminae (above: stratum oriens; below: stratum radiatum). <a href="https://cloudflare.jove.com/files/ftp_upload/56282/56282fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Metabolic Support of Excised, Living Brain Tissues During Magnetic Resonance Microscopy Acquisition at JoVE.com

Metabolic Support of Excised, Living Brain Tissues During Magnetic Resonance Microscopy Acquisition

The current protocol describes a method by which users can maintain viability of acute hippocampal and cortical slice preparations during the collection of magnetic resonance microscopy data.

This protocol describes the procedures necessary to support normal metabolic functions of acute brain slice preparations during the collection of magnetic ...

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5.9K Views.  University of Florida. The current protocol describes a method by which users can maintain viability of acute hippocampal and cortical slice preparations during the collection of magnetic resonance microscopy data.

Video: Metabolic Support of Excised, Living Brain Tissues During Magnetic Resonance Microscopy Acquisition

Magnetic Resonance Elastography Methodology for the Evaluation of Tissue Engineered Construct Growth

Traditional mechanical testing often results in the destruction of the sample, and in the case of long term tissue engineered construct studies, the use of destructive assessment is not acceptable. A proposed alternative is the use of an imaging process called magnetic resonance elastography. Elastography is a nondestructive method for determining the engineered outcome by measuring local mechanical property values (i.e., complex shear modulus), which are essential markers for identifying the structure and functionality of a tissue. As a noninvasive means for evaluation, the monitoring of engineered constructs with imaging modalities such as magnetic resonance imaging (MRI) has seen increasing interest in the past decade1. For example, the magnetic resonance (MR) techniques of diffusion and relaxometry have been able to characterize the changes in chemical and physical properties during engineered tissue development2. The method proposed in the following protocol uses microscopic magnetic resonance elastography (&mu;MRE) as a noninvasive MR based technique for measuring the mechanical properties of small soft tissues3. MRE is achieved by coupling a sonic mechanical actuator with the tissue of interest and recording the shear wave propagation with an MR scanner4. Recently, &mu;MRE has been applied in tissue engineering to acquire essential growth information that is traditionally measured using destructive mechanical macroscopic techniques5. In the following procedure, elastography is achieved through the imaging of engineered constructs with a modified Hahn spin-echo sequence coupled with a mechanical actuator. As shown in Figure 1, the modified sequence synchronizes image acquisition with the transmission of external shear waves; subsequently, the motion is sensitized through the use of oscillating bipolar pairs. Following collection of images with positive and negative motion sensitization, complex division of the data produce a shear wave image. Then, the image is assessed using an inversion algorithm to generate a shear stiffness map6. The resulting measurements at each voxel have been shown to strongly correlate (R2&gt;0.9914) with data collected using dynamic mechanical analysis7. In this study, elastography is integrated into the tissue development process for monitoring human mesenchymal stem cell (hMSC) differentiation into adipogenic and osteogenic constructs as shown in Figure 2.

The procedure demonstrates the methodology of magnetic resonance elastography for monitoring the engineered outcome of adipose and osteogenic tissue engineered constructs through noninvasive local assessment of the mechanical properties using microscopic magnetic resonance elastography (&mu;MRE).

Traditional mechanical testing often results in the destruction of the sample, and in the case of long term tissue engineered construct studies, the use ...

Magnetically-Assisted Remote Controlled Microcatheter Tip Deflection under Magnetic Resonance Imaging

X-ray fluoroscopy-guided endovascular procedures have several significant limitations, including difficult catheter navigation and use of ionizing radiation, which can potentially be overcome using a magnetically steerable catheter under MR guidance.
	The main goal of this work is to develop a microcatheter whose tip can be remotely controlled using the magnetic field of the MR scanner. This protocol aims to describe the procedures for applying current to the microcoil-tipped microcatheter to produce consistent and controllable deflections.
	A microcoil was fabricated using laser lathe lithography onto a polyimide-tipped endovascular catheter. In vitro testing was performed in a waterbath and vessel phantom under the guidance of a 1.5-T MR system using steady-state free precession (SSFP) sequencing. Various amounts of current were applied to the coils of the microcatheter to produce measureable tip deflections and navigate in vascular phantoms.
	The development of this device provides a platform for future testing and opportunity to revolutionize the endovascular interventional MRI environment.

Current applied to an endovascular microcatheter with microcoil tip made by laser lathe lithography can achieve controllable deflections under magnetic resonance (MR) guidance, which may improve speed and efficacy of navigation of vasculature during various endovascular procedures.

X-ray fluoroscopy-guided  endovascular procedures have several significant limitations, including  difficult catheter navigation and use of ionizing ...

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Mechanical properties of cells and extracellular matrix (ECM) play important roles in many biological processes including stem cell differentiation, tumor formation, and wound healing. Changes in stiffness of cells and ECM are often signs of changes in cell physiology or diseases in tissues. Hence, cell stiffness is an index to evaluate the status of cell cultures. Among the multitude of methods applied to measure the stiffness of cells and tissues, micro-indentation using an Atomic Force Microscope (AFM) provides a way to reliably measure the stiffness of living cells. This method has been widely applied to characterize the micro-scale stiffness for a variety of materials ranging from metal surfaces to soft biological tissues and cells. The basic principle of this method is to indent a cell with an AFM tip of selected geometry and measure the applied force from the bending of the AFM cantilever. Fitting the force-indentation curve to the Hertz model for the corresponding tip geometry can give quantitative measurements of material stiffness. This paper demonstrates the procedure to characterize the stiffness of living cells using AFM. Key steps including the process of AFM calibration, force-curve acquisition, and data analysis using a MATLAB routine are demonstrated. Limitations of this method are also discussed.

This paper demonstrates a protocol to characterize the mechanical properties of living cells by means of microindentation using an Atomic Force Microscope (AFM).

Mechanical  properties of cells and extracellular matrix (ECM) play important roles in many  biological processes including stem cell differentiation, ...

FIBS-enabled Noninvasive Metabolic Profiling

In the era of computational biology, new high throughput experimental systems are necessary in order to populate and refine models so that they can be validated for predictive purposes. Ideally such systems would be low volume, which precludes sampling and destructive analyses when time course data are to be obtained. What is needed is an in situ monitoring tool which can report the necessary information in real-time and noninvasively. An interesting option is the use of fluorescent, protein-based in vivo biological sensors as reporters of intracellular concentrations. One particular class of in vivo biosensors that has found applications in metabolite quantification is based on F&#246;rster Resonance Energy Transfer (FRET) between two fluorescent proteins connected by a ligand binding domain. FRET integrated biological sensors (FIBS) are constitutively produced within the cell line, they have fast response times and their spectral characteristics change based on the concentration of metabolite within the cell. In this paper, the method for constructing Chinese hamster ovary (CHO) cell lines that constitutively express a FIBS for glucose and glutamine and calibrating the FIBS in vivo in batch cell culture in order to enable future quantification of intracellular metabolite concentration is described. Data from fed-batch CHO cell cultures demonstrates that the FIBS was able in each case to detect the resulting change in the intracellular concentration. Using the fluorescent signal from the FIBS and the previously constructed calibration curve, the intracellular concentration was accurately determined as confirmed by an independent enzymatic assay.

A description of how to calibrate F&#246;rster Resonance Energy Transfer integrated biological sensors (FIBS) for in situ metabolic profiling is presented.&#160;The FIBS can be used to measure intracellular levels of metabolites noninvasively aiding in the development of metabolic models and high throughput screening of bioprocess conditions.

In the era of computational biology, new high throughput experimental systems are necessary in order to populate and refine models so that they can be ...

Custom-designed Laser-based Heating Apparatus for Triggered Release of Cisplatin from Thermosensitive Liposomes with Magnetic Resonance Image Guidance

Liposomes have been employed as drug delivery systems to target solid tumors through exploitation of the enhanced permeability and retention (EPR) effect resulting in significant reductions in systemic toxicity. Nonetheless, insufficient release of encapsulated drug from liposomes has limited their clinical efficacy. Temperature-sensitive liposomes have been engineered to provide site-specific release of drug in order to overcome the problem of limited tumor drug bioavailability. Our lab has designed and developed a heat-activated thermosensitive liposome formulation of cisplatin (CDDP), known as HTLC, to provide triggered release of CDDP at solid tumors. Heat-activated delivery in vivo was achieved in murine models using a custom-built laser-based heating apparatus that provides a conformal heating pattern at the tumor site as confirmed by MR thermometry (MRT). A fiber optic temperature monitoring device was used to measure the temperature in real-time during the entire heating period with online adjustment of heat delivery by alternating the laser power. Drug delivery was optimized under magnetic resonance (MR) image guidance by co-encapsulation of an MR contrast agent (i.e.,&#160;gadoteridol) along with CDDP into the thermosensitive liposomes as a means to validate the heating protocol and to assess tumor accumulation. The heating protocol consisted of a preheating period of 5 min prior to administration of HTLC and 20 min heating post-injection. This heating protocol resulted in effective release of the encapsulated agents with the highest MR signal change observed in the heated tumor in comparison to the unheated tumor and muscle. This study demonstrated the successful application of the laser-based heating apparatus for preclinical thermosensitive liposome development and the importance of MR-guided validation of the heating protocol for optimization of drug delivery.

AN MRI-compatible&#160;custom-designed laser-based heating apparatus has been developed to provide local heating of subcutaneous tumors in order to activate release of agents from thermosensitive liposomes specifically at the tumor region.

Liposomes have been employed as drug delivery systems to target solid tumors through exploitation of the enhanced permeability and retention (EPR) effect ...

Rapid Scan Electron Paramagnetic Resonance Opens New Avenues for Imaging Physiologically Important Parameters In Vivo

We demonstrate a superior method of 2D spectral-spatial imaging of stable radical reporter molecules at 250 MHz using rapid-scan electron-paramagnetic-resonance (RS-EPR), which can provide quantitative information under in vivo conditions on oxygen concentration, pH, redox status and concentration of signaling molecules (i.e., OH&#8226;, NO&#8226;). The RS-EPR technique has a higher sensitivity, improved spatial resolution (1 mm), and shorter acquisition time in comparison to the standard continuous wave (CW) technique. A variety of phantom configurations have been tested, with spatial resolution varying from 1 to 6 mm, and spectral width of the reporter molecules ranging from 16 &#181;T (160 mG) to 5 mT (50 G). A cross-loop bimodal resonator decouples excitation and detection, reducing the noise, while the rapid scan effect allows more power to be input to the spin system before saturation, increasing the EPR signal. This leads to a substantially higher signal-to-noise ratio than in conventional CW EPR experiments.

Rapid Scan Electron Paramagnetic Resonance Opens New Avenues for Imaging Physiologically Important Parameters In Vivo

A new electron paramagnetic resonance (EPR) method, rapid scan EPR (RS-EPR), is demonstrated for 2D spectral spatial imaging which is superior to the traditional continuous wave (CW) technique and opens new venues for in vivo imaging. Results are demonstrated at 250 MHz, but the technique is applicable at any frequency.

We demonstrate a superior method of 2D spectral-spatial imaging of stable radical reporter molecules at 250 MHz using rapid-scan ...

Automated Slide Scanning and Segmentation in Fluorescently-labeled Tissues Using a Widefield High-content Analysis System

Automated slide scanning and segmentation of fluorescently-labeled tissues is the most efficient way to analyze whole slides or large tissue sections. Unfortunately, many researchers spend large amounts of time and resources developing and optimizing workflows that are only relevant to their own experiments. In this article, we describe a protocol that can be used by those with access to a widefield high-content analysis system (WHCAS) to image any slide-mounted tissue, with options for customization within pre-built modules found in the associated software. Not originally intended for slide scanning, the steps detailed in this article make it possible to acquire slide scanning images in the WHCAS which can be imported into the associated software. In this example, the automated segmentation of brain tumor slides is demonstrated, but the automated segmentation of any fluorescently-labeled nuclear or cytoplasmic marker is possible. Furthermore, there are a variety of other quantitative software modules including assays for protein localization/translocation, cellular proliferation/viability/apoptosis, and angiogenesis that can be run. This technique will save researchers time and effort and create an automated protocol for slide analysis.

Here we describe a protocol for the automated segmentation of fluorescently labeled tissues on slides using a widefield high-content analysis system (WHCAS). This protocol has wide-ranging applications in any field which involves the quantitation of fluorescent markers in biological tissues including the biological sciences, medical engineering, and health sciences.

Automated slide scanning and segmentation of fluorescently-labeled tissues is the most efficient way to analyze whole slides or large tissue sections. ...

Controlled Strain of 3D Hydrogels under Live Microscopy Imaging

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized in vitro stretching methods. Various available systems use 2D substrate-based stretchers, while the accessibility of 3D techniques to strain soft hydrogels, is more restricted. Here, we describe a method that allows external stretching of soft hydrogels from their circumference, using an elastic silicone strip as the sample carrier. The stretching system utilized in this protocol is constructed from 3D-printed parts and low-cost electronics, making it simple and easy to replicate in other labs. The experimental process begins with polymerizing thick (&#62;100 &#956;m) soft fibrin hydrogels (Elastic Modulus of ~100 Pa) in a cut-out at the center of a silicone strip. Silicone-gel constructs are then attached to the printed-stretching device and placed on the confocal microscope stage. Under live microscopy the stretching device is activated, and the gels are imaged at various stretch magnitudes. Image processing is then used to quantify the resulting gel deformations, demonstrating relatively homogenous strains and fiber alignment throughout the gel&#8217;s 3D thickness (Z-axis). Advantages of this method include the ability to strain extremely soft hydrogels in 3D while executing in situ microscopy, and the freedom to manipulate the geometry and size of the sample according to the user&#8217;s needs. Additionally, with proper adaptation, this method can be used to stretch other types of hydrogels (e.g., collagen, polyacrylamide or polyethylene glycol) and can allow for analysis of cells and tissue response to external forces under more biomimetic 3D conditions.

The presented method involves uniaxial stretching of 3D soft hydrogels embedded in silicone rubber while allowing live confocal microscopy. Characterization of the external and internal hydrogel strains as well as fiber alignment are demonstrated. The device and protocol developed can assess the response of cells to various strain regimes.

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized ...

Electric and Magnetic Field Devices for Stimulation of Biological Tissues

Electric fields (EFs) and magnetic fields (MFs) have been widely used by tissue engineering to improve cell dynamics such as proliferation, migration, differentiation, morphology, and molecular synthesis. However, variables such stimuli strength and stimulation times need to be considered when stimulating either cells, tissues or scaffolds. Given that EFs and MFs vary according to cellular response, it remains unclear how to build devices that generate adequate biophysical stimuli to stimulate biological samples. In fact, there is a lack of evidence regarding the calculation and distribution when biophysical stimuli are applied. This protocol is focused on the design and manufacture of devices to generate EFs and MFs and implementation of a computational methodology to predict biophysical stimuli distribution inside and outside of biological samples. The EF device was composed of two parallel stainless-steel electrodes located at the top and bottom of biological cultures. Electrodes were connected to an oscillator to generate voltages (50, 100, 150 and 200 Vp-p) at 60 kHz. The MF device was composed of a coil, which was energized with a transformer to generate a current (1 A) and voltage (6 V) at 60 Hz. A polymethyl methacrylate support was built to locate the biological cultures in the middle of the coil. The computational simulation elucidated the homogeneous distribution of EFs and MFs inside and outside of biological tissues. This computational model is a promising tool that can modify parameters such as voltages, frequencies, tissue morphologies, well plate types, electrodes and coil size to estimate the EFs and MFs to achieve a cellular response.

This protocol describes the step-by-step process to build both electrical and magnetic stimulators used to stimulate biological tissues. The protocol includes a guideline to simulate computationally electric and magnetic fields and manufacture of stimulator devices.

Electric fields (EFs) and magnetic fields (MFs) have been widely used by tissue engineering to improve cell dynamics such as proliferation, migration, ...

Quantification of Mouse Heart Left Ventricular Function, Myocardial Strain, and Hemodynamic Forces by Cardiovascular Magnetic Resonance Imaging

Mouse models have contributed significantly to understanding genetic and physiological factors involved in healthy cardiac function, how perturbations result in pathology, and how myocardial diseases may be treated. Cardiovascular magnetic resonance imaging (CMR) has become an indispensable tool for a comprehensive in vivo assessment of cardiac anatomy and function. This protocol shows detailed measurements of mouse heart left ventricular function, myocardial strain, and hemodynamic forces using 7-Tesla CMR. First, animal preparation and positioning in the scanner are demonstrated. Survey scans are performed for planning imaging slices in various short- and long-axis views. A series of prospective ECG-triggered short-axis (SA) movies (or CINE images) are acquired covering the heart from apex to base, capturing end-systolic and end-diastolic phases. Subsequently, single-slice, retrospectively gated CINE images are acquired in a midventricular SA view, and in 2-, 3-, and 4-chamber views, to be reconstructed into high-temporal resolution CINE images using custom-built and open-source software. CINE images are subsequently analyzed using dedicated CMR image analysis software.
Delineating endomyocardial and epicardial borders in SA end-systolic and end-diastolic CINE images allows for the calculation of end-systolic and end-diastolic volumes, ejection fraction, and cardiac output. The midventricular SA CINE images are delineated for all cardiac time frames to extract a detailed volume-time curve. Its time derivative allows for the calculation of the diastolic function as the ratio of the early filling and atrial contraction waves. Finally, left ventricular endocardial walls in the 2-, 3-, and 4-chamber views are delineated using feature-tracking, from which longitudinal myocardial strain parameters and left ventricular hemodynamic forces are calculated. In conclusion, this protocol provides detailed in vivo quantification of the mouse cardiac parameters, which can be used to study temporal alterations in cardiac function in various mouse models of heart disease.

This study describes a comprehensive cardiovascular magnetic resonance imaging (CMR) protocol to quantify the left ventricular functional parameters of the mouse heart. The protocol describes the acquisition, post-processing, and analysis of the CMR images as well as assessment of different cardiac functional parameters.

Mouse models have contributed significantly to understanding genetic and physiological factors involved in healthy cardiac function, how perturbations ...