This work was supported by grants from the Cancer Research Prevention &#38; Research Institute of Texas [RR150076], the National Science Foundation [1751174]; the National Institutes of Health [NS-111236], the Welch Foundation [1-1923-20170325]; the Lupe Murchison Foundation, the Ted Nash Long Life Foundation and the Kinship Foundation (Searle Scholars Program) to K.K.F.

1. Culture and cryoprotection of mammalian cells
<ol>
	<li>Culture and harvesting mammalian cells
	<ol>
		<li>Thaw an aliquot of frozen human embryonic kidney cells (HEK 293).</li>
		<li>Culture HEK 293 cells in growth media (e.g., DMEM with 10% fetal bovine serum and 1% Pen-Strep) at 37 &#176;C with 5% CO2 in 100 mm plates for two to three passages (7-10 days).</li>
		<li>Split the cells and culture in a 150 mm plate until cells attain 90-95% confluency. 
		NOTE: A 150 mm plate at &#62; 90% confluency will be sufficient to fill two sapphire rotors with a diameter of 3.2 mm.</li>
		<li>Harvest cells using 4 mL of trypsin (see Table of Materials) and 10 mL of media. Transfer the suspension to a sterile 15 mL conical and centrifuge at 673 x g (1000 rpm) for 5 min at room temperature. Remove the supernatant.</li>
		<li>Wash the cell pellet with phosphate buffered saline (PBS) (pH 7.4, &#8722;CaCl2, &#8722;MgCl2).</li>
	</ol>
	</li>
	<li>Cryoprotection of cells
	<ol>
		<li>Collect a 50 &#181;L cell pellet in a microcentrifuge tube. Prepare a mixture of 50 &#181;L of PBS and 18 &#181;L of glycerol in a separate tube.</li>
		<li>Gently mix the 50 &#181;L cell pellet with 68 &#181;L of glycerol-PBS mixture by adding the glycerol-PBS mixture to the top of the pellet and resuspending the pellet by gently tapping the side of the tube until no clumps remain. 
		NOTE: Gentle pipetting can also be used to resuspend the cell, however, make sure that cellular integrity is not compromised.</li>
	</ol>
	</li>
</ol>
2. Cryopreservation of mammalian cells in an NMR rotor
<ol>
	<li>Transfer of cells into a 3.2 mm sapphire rotor.
	<ol>
		<li>To make a funnel, cut a 200 &#181;L pipette tip and insert the narrow end of the cut pipette tip into the 3.2 mm rotor.</li>
		<li>Transfer the cells into the funnel sitting on the rotor. Place the rotor together with the funnel in a microcentrifuge tube and pellet the cells into the bottom of the rotor by centrifugation at 673 x g for 2-3 min at room temperature.</li>
		<li>Remove the supernatant and any excess sample from the rotor. Repeat these two steps until the rotor is fully packed with the cells. 
		NOTE: Optionally, determine the cellular viability of the sample before freezing. Resuspend 10 &#181;L of the excess cell pellet in 100 &#181;L of FBS-free DMEM. Mix 10 &#181;L of the suspension with 0.4% trypan blue solution and immediately assess viability using an automated cell counter. Use only FBS free media to dilute cells as serum interferes with trypan blue staining.</li>
		<li>Seal the rotor with a silicon plug using commercially available packing tool.</li>
		<li>Close the rotor with a ceramic drive tip by pressing it vertically downward. Avoid touching the delicate fins on the side of the drive tip.</li>
		<li>Mark half of the bottom edge of the sapphire rotor with a silver permanent marker and the other half of the bottom edge of the rotor with a black permanent marker to allow accurate monitoring of the spinning of the rotor inside of the spectrometer. 
		NOTE: Imperfections in the marking of the rotor will prevent accurate counting of the spinning frequency, resulting in the failure to achieve stable spinning. Because markers will not write on a frozen rotor and rotor warming compromises sample integrity, marking the rotor before freezing is a critical step.</li>
	</ol>
	</li>
	<li>Cryopreservation of cells inside a 3.2 mm sapphire rotor.
	<ol>
		<li>Place a cushion made by a piece of tissue or paper towel under the lid and at the bottom of the cryogenic vial (see Table of Materials). 
		NOTE: The tissue paper protects the rotor marking from damage incurred by bumping against the sides of cryogenic vials.</li>
		<li>Place the 3.2 mm sapphire rotor into the cryogenic vial padded with the tissue paper with marked end facing the bottom of the cryogenic vial.</li>
		<li>Slow freeze the rotor by placing the cryogenic vial into the controlled rate (-1 &#176;C/min) cooling container and place the container in -80 &#176;C freezer for a minimum of 3 h.</li>
		<li>Transfer the cryogenic vial containing the frozen rotor to liquid nitrogen storage.</li>
	</ol>
	</li>
</ol>
3. Cryogenic transfer of a frozen sample into the NMR spectrometer
<ol>
	<li>Transport the frozen sample to the NMR facility.
	<ol>
		<li>Transfer the cryogenic vial containing the frozen rotor to a small dewar filled with liquid nitrogen for transport to the NMR facility.</li>
	</ol>
	</li>
	<li>Transfer of frozen rotor to the liquid nitrogen bath.
	<ol>
		<li>Fill a dry, thermally insulated wide mouth foam dewar with 500 mL &#8211; 1 L of liquid nitrogen.</li>
		<li>Transfer the rotor from the cryogenic vial into the wide mouth foam dewar filled with liquid nitrogen.
		<ol>
			<li>Take the cryogenic vial from the transfer dewar in the hand and hold it just above the surface of the liquid nitrogen to protect it from the atmosphere.</li>
			<li>While holding the cryogenic vial with the mouth pointing slightly downwards, unscrew the cap and let the rotor slide into the liquid nitrogen bath. 
			NOTE: Once the cap is unscrewed, the rotor must fall from the cryogenic vial into the liquid nitrogen bath quickly (under 1 second) to prevent condensation from collecting on the rotor. The evaporation of liquid nitrogen forms a &#8220;nitrogen cloud&#8221; in the wide mouth foam dewar and prevents condensation on the rotor before it is submerged into the liquid nitrogen. Longer exposure of the rotor to air can lead to condensation of moisture on rotor walls which will re-condense into ice.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Cryogenic transfer of rotor to NMR sample catcher.
	<ol>
		<li>Prechill a 1.5 mL microcentrifuge tube by submerging it in the liquid nitrogen bath. Do not close the tube. 
		NOTE: Inspect the rotor before transfer into the microcentrifuge tube. Using tweezers, hold the rotor just below the surface of the liquid nitrogen and check that the rotor markings are intact, that no ice deposits have formed on its walls and that the drive tip is intact. Ice crystal deposits on the rotor walls appear as white powder. Be careful to always hold the rotor by its body and not by the drive tip. Do not scratch the marking off the rotor with the tweezer-tips.</li>
		<li>Under the surface of the liquid nitrogen bath, use tweezers to transfer the rotor into the microcentrifuge tube with the drive tip facing the bottom of the microcentrifuge tube and the markings facing the opening of the tube. 
		NOTE: Always dry tweezers before submerging in liquid nitrogen or touching the rotor.</li>
		<li>Using tweezers, hold the tube containing the rotor under liquid nitrogen by its neck.</li>
		<li>With a second pair of tweezers in hand, be prepared to&#160;submerge the NMR sample catcher in the liquid nitrogen bath and hold it so that it is inclined at an acute angle with respect to the microcentrifuge tube. 
		NOTE: Minimize the time that the sample catcher is in contact with the liquid nitrogen to avoid freezing the O-ring. If the O-ring freezes, it will be very difficult to insert the catcher into the spectrometer.</li>
	</ol>
	</li>
	<li>Cryogenic rotor transfer from the NMR sample catcher to NMR spectrometer 
	NOTE: This step requires two people, one to operate the cryocabinet and one to transfer the sample from the liquid nitrogen into the probe.
	<ol>
		<li>Place the cryocabinet in ejection mode by pressing &#8216;EJECT&#8217; on the cabinet. 
		NOTE: The ejection mode purges the dry and cold nitrogen gas flow at high pressure from the probe to the atmosphere in order to prevent the entrance of atmospheric moisture.</li>
		<li>Transfer the rotor into the NMR sample catcher.
		<ol>
			<li>Insert the open end of the NMR sample catcher into the microcentrifuge tube while still under the surface of the liquid nitrogen.</li>
			<li>Lift up both the microcentrifuge tube and NMR sample catcher to allow the rotor to fall into the sample catcher. Shake the NMR sample catcher and microcentrifuge tube in case the rotor is stuck on the rim of the sample catcher.</li>
			<li>Leave the empty microcentrifuge tube on top of the NMR sample catcher to shield the rotor from air.</li>
		</ol>
		</li>
		<li>Remove the other empty NMR sample catcher from the probe and lay it on the floor.</li>
		<li>Transfer the NMR sample catcher containing the rotor to your free hand, remove the microcentrifuge tube and insert it immediately into the probe. 
		NOTE: If the O-ring freezes, it will be difficult to tighten the sample catcher. Keep applying force until it slides into place.</li>
		<li>Signal the person operating the cryocabinet to &#8216;STOP EJECT&#8217; and &#8216;INSERT&#8217;. 
		NOTE: The insertion mode guides the rotor from the sample catcher into the probe.</li>
		<li>Spin up the sample to the desired spinning rate (e.g., 12 kHz) by adjusting the bearing and driving flow pressure controlled by the cryocabinet. (e.g., Immediately increase the bearing gas to ~200 mBar and the drive gas to 10 mBar. Once the sample spins, increase the bearing gas to 1000 mBar and drive gas to 200 mBar. As spinning stabilizes, increase the bearing to 2400 mBar and then increase drive gas from 200 mBar to 1700 mBar over several minutes. VT cooling gas is constant at ~1070 L/h.) 
		NOTE: When lifting the microcentrifuge tube and NMR sample catcher out of the liquid nitrogen bath, make sure that microcentrifuge tube has enough liquid nitrogen inside it to surround the rotor. Minimize the time between transferring the rotor into the NMR sample catcher and inserting the NMR sample catcher into the spectrometer. All the steps in 3.4 should be completed within 30 s.</li>
	</ol>
	</li>
</ol>
4. Cryogenic removal of the sample from the NMR spectrometer
<ol>
	<li>Preparation of the liquid nitrogen bath and the cryogenic vial
	<ol>
		<li>Pour 500 mL - 1 L of liquid nitrogen into the wide mouth foam dewar and place the bath under the spectrometer.</li>
		<li>Pre-cool the cryogenic vial. Submerge the empty cryogenic vial containing a piece of tissue paper in the liquid nitrogen bath.</li>
	</ol>
	</li>
	<li>Cryogenic transfer of rotor from probe to cryogenic vial
	<ol>
		<li>Reduce spinning rate to 0 kHz by ramping down the driving and bearing gas flow and eject the rotor by switching to the ejection mode.</li>
		<li>Keep the ejection mode on, remove the sample catcher from the probe, drop rotor directly into the wide mouth foam dewar containing liquid nitrogen.</li>
		<li>Using pre-chilled tweezers, transfer the rotor into pre-chilled cryogenic vial under the surface of the liquid nitrogen.</li>
		<li>Cap the cryogenic vial. Prechill the cryogenic vial cap by dipping it into liquid nitrogen. Remove the cryogenic vial containing the rotor and liquid nitrogen from the bath and cap the tube with prechilled cap. Do not tighten the cap so that vaporizing nitrogen can be safely released.</li>
		<li>Re-submerge the cryogenic vial in liquid nitrogen. The sample can be transferred to longer term liquid nitrogen storage or unpacked immediately for further analysis.</li>
	</ol>
	</li>
</ol>
5. Unpacking rotor and viability measurements
<ol>
	<li>Unpacking rotor and measuring viability
	<ol>
		<li>Pre-warm serum free media (DMEM) or PBS to 37 &#176;C.</li>
		<li>Remove rotor from liquid nitrogen. Remove drive tip and silicon plug. 
		NOTE: Sapphire is an excellent heat conductor. Avoid touching the rotor with your fingers because heat transfer can cause local freeze-thaw events that compromise cellular viability.</li>
	</ol>
	</li>
	<li>Measuring viability
	<ol>
		<li>Add 20 &#181;L of warm media to the frozen cell pellet in the rotor and resuspend cells. Remove suspension with a pipette and mix the suspension with 100 &#181;L of media.</li>
		<li>Remove 10 &#181;L of cell suspension and mix with equal volume of 0.4% trypan blue solution (v/v). Incubate at room temperature for 30 s to 1 min.</li>
		<li>Measure viability using automated cell counter.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The cryogenic transfer of frozen samples into an NMR spectrometer is successful in preserving the viability of frozen mammalian cells through the NMR data acquisition. The success of this methodology is demonstrated in pre and post MAS NMR viability measurements. This approach is successful and generalizable to any system where temperature fluctuations may compromise sample integrity. The currently presented protocol is performed with the HEK 293 cell line. Because cryopreservation conditions for many mammalian cell lines are very similar, it is likely that the conditions reported here are translatable to other cellular systems; however, they may require further optimization of cryoprotectants, sample volumes, and freezing rates to attain the same results.
This methodology can be improved upon by unpacking the rotor faster post NMR experiment. This step is currently sub optimal and its execution affects the viability of the cells. Before the cells can be resuspended in media, the drive tip and silicon plug must be removed from the rotor. The sample thaws unevenly when the rotor is held during removal of the drive tip and the silicon plug so shorter rotor handling times result in higher viabilities. The development of rotor holders or other tools to facilitate uniform thawing and quick removal of the drive tip and the silicon plug would aid in making the post-NMR assessment of viability more accurate.
The approach to cryogenic sample loading described in this article is limited to NMR probes that support sample insertion and ejection with the probe in place in the bore of the NMR magnet. While external sample insertion and ejection is standard for commercial DNP systems, custom probes do not always have this option. Also, this approach to cryogenic sample loading may require some modification for NMR probes built to be compatible with different sized rotors. This protocol has been optimized for 3.2 mm rotors and may require modification if the outer diameter of the sample catcher exceeds the inner diameter of a microcentrifuge tube (e.g., step 3.4.2).
With the application of DNP to MAS NMR, it is now possible to detect proteins and other biomolecules at endogenous physiological concentrations24,25,26. This opens the possibility of studying biomolecules within their native environments. Maintenance of cellular integrity and viability throughout the experiment is likely to be critical in connecting the experimental outcomes of the spectroscopy to biological phenomena. Uncontrolled freezing of samples containing purified proteins or cellular lysates does not typically compromise sample quality7,27, although there are some indications that freezing rate may be an important variable even in purified systems28. However, samples of mammalian cells need to be frozen at controlled rate if preserving cellular intactness and viability is important for the interpretation. Here we present a protocol for freezing and transferring frozen samples of mammalian cells into a pre-cooled DNP MAS NMR instrument that avoids potentially damaging temperature fluctuations and supports measurement on viable cells.

The introduction of dynamic nuclear polarization for magic angle spinning nuclear magnetic resonance spectroscopy can increase the sensitivity of MAS NMR by several orders of magnitude. This has enabled detection of biomolecules at or near their physiological concentrations. DNP can and does provide the sensitivity required to detect an isotopically labeled protein at endogenous (~1 &#181;M) concentrations in complex biological environment1. Because there are well-established protocols to introduce isotopically labeled molecules into unlabeled mammalian cells without affecting their viability, this opens the possibility of studying isotopically enriched biomolecules at their endogenous levels in their native environment. Moreover, because DNP enhancements are more efficient at lower temperatures2,3,4, the experimental temperatures for DNP MAS NMR align neatly with those required for long-term storage of viable mammalian cells5. However, the conventional method of transferring a sample into a DNP MAS NMR spectrometer subjects it to temperature fluctuation rates that rupture mammalian cells.
MAS NMR experiments require that the sample be rotated about the magic angle at frequencies equal to or greater than the magnitude of the anisotropic interaction for it to be averaged to zero, typically at least 4 kHz and often much higher6,7,8,9. Samples are, therefore, packed into rotors that have a finned tip that is used to drive the rotation of the rotor by a stream of gas and have a mark at the other end so the rotation frequency can be monitored by a tachometer. Sample transfer for most MAS NMR instruments is accomplished by injecting the rotor from the exterior of the instrument into the stator at the end of the NMR probe with a stream of dry air or nitrogen gas. After the rotor reaches the stator, which holds the rotor at the magic angle, sample rotation is propelled by an air turbine mechanism. Separate streams of gas support, propel and control the temperature of the rotor. Inserting a rotor into the NMR spectrometer and achieving stable MAS spinning requires finely machined drive tips and tight control of the temperature and pressure of the separate streams of gas. Despite these technical demands, insertion and achieving stable MAS are largely automated for commercial MAS NMR probes for room-temperature applications.
However, the situation is more complicated for low temperature applications. Samples for low temperature applications are typically inserted into the spectrometer at room temperature and frozen in the stator. In the first minute, the sample temperature decreases quickly (&#62; &#8722;100 &#176;C/min) and the system temperature requires several minutes to equilibrate. Because of the interplay of temperature and pressure, insertion and approaching of the desired MAS are often handled manually for low-temperature applications. Despite the requirement for manual intervention, freezing the rotor inside of the instrument is beneficial because it minimizes the introduction of water and condensation into the probe, which is critical for successful spinning. Not only can condensation and ice build-up from ambient moisture block gas lines, condensation, or frost on the rotor itself can mechanically prevent MAS. Thus, samples for low temperature MAS NMR are typically frozen inside of the instrument at rates that exceed -100 &#176;C/min.
Mammalian cells can retain their integrity through a freeze-thaw cycle if the cooling is slow5,10,11,12, at a rate equal or slower than 1 &#176;C/min. Alternatively cells also retain their integrity if the cooling rate is ultra-fast13,14,15, at a rate faster than 104 &#176;C/min. Rates intermediate to these two extremes rupture and kill mammalian cells due to ice crystal formation both inside and outside the cells, even in the presence of cryoprotective agents16. The sample cooling rates for a room temperature rotor inside a pre-cooled probe fall between these two extremes, thus to study cryogenically preserved intact viable mammalian cells, samples must be frozen before transfer into the instrument and transferred into the instrument without temperature fluctuations that could damage the sample or accumulation of frost on the rotor that could prevent the rotor from spinning. The protocol describes a method for frost-free, pre-cooled rotor insertion into a cryogenic MAS NMR system for the study of cryogenically preserved intact viable mammalian cell samples. The cryogenic sample transfer described here was developed for NMR characterization of viable intact cells. However, it is applicable to any system where temperature fluctuations may compromise sample integrity. This includes any variety of complex systems, such as freeze quenched reactions for chemical and structural characterization of trapped reaction intermediates17,18, enzymology19,20 or protein folding21,22.

<table><tbody><tr><td>0.4% Trypan blue stain</td><td>Invitrogen</td><td>T10282</td><td></td></tr><tr><td>100 mm cell culture dish</td><td>Thomas Scientific</td><td>430167</td><td></td></tr><tr><td>150 mm cell culture dish</td><td>Nunc</td><td>157150</td><td></td></tr><tr><td>3.2 mm sapphire rotor</td><td>Bruker</td><td></td><td></td></tr><tr><td>45 &amp;deg; angled forceps</td><td>Hampton Research</td><td>HR4-859</td><td></td></tr><tr><td>AMUPol</td><td>Cortecnet</td><td>C010P005</td><td></td></tr><tr><td>Black and Silver marker</td><td>Sharpie</td><td></td><td></td></tr><tr><td>Cap removal tool</td><td>Bruker</td><td>HZ16913</td><td></td></tr><tr><td>Cell culture grade water</td><td>HyClone</td><td>SH30529.03</td><td></td></tr><tr><td>Ceramic cap</td><td>Cortecnet</td><td>HZ12372</td><td></td></tr><tr><td>CoolCell</td><td>Corning/ Biocision</td><td>UX-04392-00 (Corning) / BCS-405 (Bioscion)</td><td></td></tr><tr><td>Countess Cell Counting Chamber Slides</td><td>Invitrogen</td><td>C10288</td><td></td></tr><tr><td>Countess II automated cell counter</td><td>Invitrogen</td><td>AMQAF1000</td><td></td></tr><tr><td>Cryogen tubes</td><td>Nalgene</td><td>03-337-7Y</td><td></td></tr><tr><td>d8-glycerol</td><td>Aldrich</td><td>447498</td><td></td></tr><tr><td>Deuterium oxide, 99.8 % atom D</td><td>Aldrich</td><td>756822-1</td><td></td></tr><tr><td>DMEM</td><td>Gibco</td><td>10569-010</td><td></td></tr><tr><td>DNP NMR system with frequency-matched gyrotron</td><td>Bruker</td><td></td><td></td></tr><tr><td>Foam dewar</td><td>Spearlab</td><td>FD-800</td><td></td></tr><tr><td>Kimwipes</td><td>Kimwipes</td><td></td><td></td></tr><tr><td>Packing tool</td><td>Bruker</td><td></td><td></td></tr><tr><td>Pen-Strep</td><td>Gibco</td><td>15140-122</td><td></td></tr><tr><td>Powdered PBS</td><td>VWR</td><td>VWRRV0780</td><td></td></tr><tr><td>Protonated PBS</td><td>Gibco</td><td>10010-023</td><td></td></tr><tr><td>Silicon plug</td><td>Bruker</td><td>B7089</td><td></td></tr><tr><td>Standard Vessel forceps</td><td></td><td></td><td></td></tr><tr><td>Tryp-L</td><td>Gibco</td><td>12605010</td><td></td></tr></tbody></table>

cryogenic sample loading into a magic angle spinning nuclear magnetic resonance spectrometer that preserves cellular viability

Dynamic nuclear polarization (DNP) can dramatically increase the sensitivity of magic angle spinning (MAS) nuclear magnetic resonance (NMR) spectroscopy. These sensitivity gains increase as temperatures decrease and are large enough to enable the study of molecules at very low concentrations at the operating temperatures (~100 K) of most commercial DNP-equipped NMR spectrometers. This leads to the possibility of in-cell structural biology on cryopreserved cells for macromolecules at their endogenous levels in their native environments. However, the freezing rates required for cellular cryopreservation are exceeded during typical sample handling for DNP MAS NMR and this results in loss of cellular integrity and viability. This article describes a detailed protocol for the preparation and cryogenic transfer of a frozen sample of mammalian cells into a MAS NMR spectrometer.

Cryogenic insertion of pre-frozen samples of mammalian cells into the NMR spectrometer supports viability throughout the NMR experiment. Schematics of cryogenic transfer of a frozen sample into a pre-cooled NMR probe is shown in Figure 1. Cellular viability and intactness can be assessed using a variety of methods. Here we used a standard dye-based measure of membrane integrity, which aligns well with other methods23. Intact cells are impermeable to trypan blue while cells with compromised membrane integrity are permeable. The number of trypan blue permeable and impermeable cells can be rapidly assessed using an automated cell counter. Using the protocol described here, the trypan blue permeability of mammalian cells after MAS NMR (i.e., at point 5.2.3) is similar to the trypan blue permeability of mammalian cells before any temperature change (i.e., point 2.1.3). However, if cells are slow frozen, then warmed to room temperature before insertion (i.e. following the protocol to point 3 before warming the rotor to room temperature before inserting into the chilled probe), cellular viability as assessed by trypan blue decreases to less than 10% of cells (Figure 2). Thus, freezing cells inside the spectrometer results in the loss of cellular membrane integrity while cryogenic insertion of frozen samples of mammalian cells supports cell viability throughout the NMR experiment.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61733/61733fig01.jpg" /> 
Figure 1: Schematic of cryogenic transfer of a frozen sample into a pre-cooled NMR probe.&#160;(A) Approach the surface of the liquid nitrogen and unscrew the top of the cryogenic vial. (B) Slide the rotor into the liquid nitrogen bath. (C) Submerge a microcentrifuge tube using tweezers and hold it in place until it cools completely. (D) Insert the rotor in the microcentrifuge with the drive tip facing the bottom of the tube. Push, rather than grab, with the tweezers. (E) Hold the microcentrifuge tube just below the surface of the liquid nitrogen bath and visually inspect the rotor to ensure that it is frost free and well-marked. (F) Hold the sample catcher at an angle above the surface. (G) Lift the sample catcher and tube out of the liquid nitrogen as soon as the sample catcher is inside the microcentrifuge tube. (H) Shake the sample catcher if the rotor is caught on the rim. (I) Remove the empty sample catcher from the probe and lay it on the ground. (J) Remove the microcentrifuge tube from the sample catcher with the rotor inside, insert, tighten the sample catcher in the probe and press &#8220;INSERT&#8221; on the control console. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61733/61733fig01large.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/61733/61733fig02.jpg" /> 
Figure 2: Cryogenic insertion of a pre-frozen sample of mammalian cells results in measurements of cellular intactness that are similar to mammalian cells samples that have never been frozen.&#160;The percentage of trypan blue impermeable cells for samples that have never been frozen (e.g. step 2.1.3) is similar to that of cells for samples after MAS NMR (e.g. step 5.2.3 with 12 kHz MAS). Slow frozen cells (e.g. step 3) that were warmed to room temperature before insertion into the NMR instrument that had been pre-cooled to 100 K had much lower percentages of intact cells after MAS NMR with 12 kHz MAS. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61733/61733fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Cryogenic Sample Loading into a Magic Angle Spinning Nuclear Magnetic Resonance Spectrometer that Preserves Cellular Viability at JoVE.com

Cryogenic Sample Loading into a Magic Angle Spinning Nuclear Magnetic Resonance Spectrometer that Preserves Cellular Viability

Presented here is a protocol for cryogenic transfer of frozen samples into the dynamic nuclear polarization (DNP) magic angle spinning (MAS) nuclear magnetic resonance (NMR) probe. The protocol includes directions for rotor storage prior to the experiment and directions for viability measurements before and after the experiment.

Dynamic nuclear polarization (DNP) can dramatically increase the sensitivity of magic angle spinning (MAS) nuclear magnetic resonance (NMR) spectroscopy. ...

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Nanyang Technological University

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Biology

3.4K Views.  University of Texas Southwestern Medical Center. Hier wordt een protocol gepresenteerd voor cryogene overdracht van bevroren monsters in de dynamische nucleaire polarisatie (DNP) magic angle spinning (MAS) nucleaire magnetische resonantie (NMR) sonde. Het protocol bevat aanwijzingen voor rotoropslag voorafgaand aan het experiment en aanwijzingen voor levensvatbaarheidsmetingen voor en na het experiment.

Video: Cryogenic Sample Loading into a Magic Angle Spinning Nuclear Magnetic Resonance Spectrometer that Preserves Cellular Viability

In Situ Detection and Single Cell Quantification of Metal Oxide Nanoparticles Using Nuclear Microprobe Analysis

Micro-analytical techniques based on chemical element imaging enable the localization and quantification of chemical composition at the cellular level. They offer new possibilities for the characterization of living systems and are particularly appropriate for detecting, localizing and quantifying the presence of metal oxide nanoparticles both in biological specimens and the environment. Indeed, these techniques all meet relevant requirements in terms of (i) sensitivity (from 1 up to 10 &#181;g.g-1 of dry mass), (ii) micrometer range spatial resolution, and (iii) multi-element detection. Given these characteristics, microbeam chemical element imaging can powerfully complement routine imaging techniques such as optical and fluorescence microscopy. This protocol describes how to perform a nuclear microprobe analysis on cultured cells (U2OS) exposed to titanium dioxide nanoparticles. Cells must grow on and be exposed directly in a specially designed sample holder used on the optical microscope and in the nuclear microprobe analysis stages. Plunge-freeze cryogenic fixation of the samples preserves both the cellular organization and the chemical element distribution. Simultaneous nuclear microprobe analysis (scanning transmission ion microscopy, Rutherford backscattering spectrometry and particle induced X-ray emission) performed on the sample provides information about the cellular density, the local distribution of the chemical elements, as well as the cellular content of nanoparticles. There is a growing need for such analytical tools within biology, especially in the emerging context of Nanotoxicology and Nanomedicine for which our comprehension of the interactions between nanoparticles and biological samples must be deepened. In particular, as nuclear microprobe analysis does not require nanoparticles to be labelled, nanoparticle abundances are quantifiable down to the individual cell level in a cell population, independently of their surface state.

In Situ Detection and Single Cell Quantification of Metal Oxide Nanoparticles Using Nuclear Microprobe Analysis

We describe a procedure for the detection of chemical elements present in situ in human cells as well as their in vitro quantification. The method is well-suited to any cell type and is particularly useful for quantitative chemical analyses in single cells following in vitro metal oxide nanoparticles exposure.

Micro-analytical techniques based on chemical element imaging enable the localization and quantification of chemical composition at the cellular level. ...

Chemistry

Cell Culture on Silicon Nitride Membranes and Cryopreparation for Synchrotron X-ray Fluorescence Nano-analysis

Very little is known about the distribution of metal ions at the subcellular level. However, those chemical elements have essential regulatory functions and their disturbed homeostasis is involved in various diseases. State-of-the-art synchrotron X-ray fluorescence nanoprobes provide the required sensitivity and spatial resolution to elucidate the two-dimensional (2D) and three-dimensional (3D) distribution and concentration of metals inside entire cells at the organelle level. This opens new exciting scientific fields of investigation on the role of metals in the physiopathology of the cell. The cellular preparation is a key and often complex procedure, particularly for basic analysis. Although X-ray fluorescence techniques are now widespread and various preparation methods have been used, very few studies have investigated the preservation of the elemental content of cells at best, and no stepwise detailed protocol for the cryopreparation of adherent cells for X-ray fluorescence nanoprobes has been released so far. This is a description of a protocol that provides the stepwise cellular preparation for fast cryofixation to enable synchrotron X-ray fluorescence nano-analysis of cells in a frozen hydrated state when a cryogenic environment and transfer is available. In case nano-analysis has to be performed at room temperature, an additional procedure for freeze-drying the cryofixed adherent cellular preparation is provided. The proposed protocols have been successfully used in previous works, most recently in studying the 2D and 3D intracellular distribution of an organometallic compound in breast cancer cells.

Presented here is a protocol for cell culture on silicon nitride membranes and plunge-freezing prior to X-ray fluorescence imaging with a synchrotron cryogenic X-ray nanoprobe. When only room temperature nano-analysis is provided, the frozen samples can be further freeze-dried. These are critical steps to obtain information on the intracellular elemental composition.

Very little is known about the distribution of metal ions at the subcellular level. However, those chemical elements have essential regulatory functions ...

Biochemistry

Biological Samples Preparation for Speciation at Cryogenic Temperature using High-Resolution  X-Ray Absorption Spectroscopy

The study of elements with X-ray absorption spectroscopy (XAS) is of particular interest when studying the role of metals in biological systems. Sample preparation is a key and often complex procedure, particularly for biological samples. Although X-ray speciation techniques are widely used, no detailed protocol has been yet disseminated for users of the technique. Further, chemical state modification is of concern, and cryo-based techniques are recommended to analyze the biological samples in their near-native hydrated state to provide the maximum preservation of chemical integrity of the cells or tissues. Here, we propose a cellular preparation protocol based on cryo-preserved samples. It is demonstrated in a high energy resolution fluorescence detected X-ray absorption spectroscopy study of selenium in cancer cells and a study of iron in phytoplankton. This protocol can be used with other biological samples and other X-ray techniques that can be damaged by irradiation.

This protocol presents a detailed procedure to prepare biological cryosamples for synchrotron-based X-ray absorption spectroscopy experiments. We describe all the steps required to optimize sample preparation and cryopreservation with examples of the protocol with cancer and phytoplankton cells. This method provides a universal standard of sample cryo-preparation.

The study of elements with X-ray absorption spectroscopy (XAS) is of particular interest when studying the role of metals in biological systems. Sample ...

Nanoscale Characterization of Liquid-Solid Interfaces by Coupling Cryo-Focused Ion Beam Milling with Scanning Electron Microscopy and Spectroscopy

Physical and chemical processes at solid-liquid interfaces play a crucial role in many natural and technological phenomena, including catalysis, solar energy and fuel generation, and electrochemical energy storage. Nanoscale characterization of such interfaces has recently been achieved using cryogenic electron microscopy, thereby providing a new path to advancing our fundamental understanding of interface processes.
This contribution provides a practical guide to mapping the structure and chemistry of solid-liquid interfaces in materials and devices using an integrated cryogenic electron microscopy approach. In this approach, we pair cryogenic sample preparation which allows stabilization of solid-liquid interfaces with cryogenic focused ion beam (cryo-FIB) milling to create cross-sections through these complex buried structures. Cryogenic scanning electron microscopy (cryo-SEM) techniques performed in a dual-beam FIB/SEM enable direct imaging as well as chemical mapping at the nanoscale. We discuss practical challenges, strategies to overcome them, as well as protocols for obtaining optimal results. While we focus in our discussion on interfaces in energy storage devices, the methods outlined are broadly applicable to a range of fields where solid-liquid interface play a key role.

Cryogenic Focused Ion Beam (FIB) and Scanning Electron Microscopy (SEM) techniques can provide key insights into the chemistry and morphology of intact solid-liquid interfaces. Methods for preparing high quality Energy Dispersive X-ray (EDX) spectroscopic maps of such interfaces are detailed, with a focus on energy storage devices.

Physical and chemical processes at solid-liquid interfaces play a crucial role in many natural and technological phenomena, including catalysis, solar ...

Engineering

High-Temperature and High-Pressure In situ Magic Angle Spinning Nuclear Magnetic Resonance Spectroscopy

Nuclear magnetic resonance (NMR) spectroscopy represents an important technique to understand the structure and bonding environments of molecules. There exists a drive to characterize materials under conditions relevant to the chemical process of interest. To address this, in situ high-temperature, high-pressure MAS NMR methods have been developed to enable the observation of chemical interactions over a range of pressures (vacuum to several hundred bar) and temperatures (well below 0 &#176;C to 250 &#176;C). Further, the chemical identity of the samples can be comprised of solids, liquids, and gases or mixtures of the three. The method incorporates all-zirconia NMR rotors (sample holder for MAS NMR) which can be sealed using a threaded cap to compress an O-ring. This rotor exhibits great chemical resistance, temperature compatibility, low NMR background, and can withstand high pressures. These combined factors enable it to be utilized in a wide range of system combinations, which in turn permit its use in diverse fields as carbon sequestration, catalysis, material science, geochemistry, and biology. The flexibility of this technique makes it an attractive option for scientists from numerous disciplines.

The molecular structures and dynamics of solids, liquids, gases, and mixtures are of critical interest to diverse scientific fields. High-temperature, high-pressure in situ MAS NMR enables detection of the chemical environment of constituents in mixed phase systems under tightly controlled chemical environments.

Nuclear magnetic resonance (NMR) spectroscopy represents an important technique to understand the structure and bonding environments of molecules. There ...

Electron Cryotomography of Bacterial Cells

While much is already known about the basic metabolism of bacterial cells, many fundamental questions are still surprisingly unanswered, including for instance how they generate and maintain specific cell shapes, establish polarity, segregate their genomes, and divide. In order to understand these phenomena, imaging technologies are needed that bridge the resolution gap between fluorescence light microscopy and higher-resolution methods such as X-ray crystallography and NMR spectroscopy.
Electron cryotomography (ECT) is an emerging technology that does just this, allowing the ultrastructure of cells to be visualized in a near-native state, in three dimensions (3D), with "macromolecular" resolution (~4nm).1, 2 In ECT, cells are imaged in a vitreous, "frozen-hydrated" state in a cryo transmission electron microscope (cryoTEM) at low temperature (&lt; -180&deg;C). For slender cells (up to ~500 nm in thickness3), intact cells are plunge-frozen within media across EM grids in cryogens such as ethane or ethane/propane mixtures. Thicker cells and biofilms can also be imaged in a vitreous state by first "high-pressure freezing" and then, "cryo-sectioning" them. A series of two-dimensional projection images are then collected through the sample as it is incrementally tilted along one or two axes. A three-dimensional reconstruction, or "tomogram" can then be calculated from the images. While ECT requires expensive instrumentation, in recent years, it has been used in a few labs to reveal the structures of various external appendages, the structures of different cell envelopes, the positions and structures of cytoskeletal filaments, and the locations and architectures of large macromolecular assemblies such as flagellar motors, internal compartments and chemoreceptor arrays.1, 2
In this video article we illustrate how to image cells with ECT, including the processes of sample preparation, data collection, tomogram reconstruction, and interpretation of the results through segmentation and in some cases correlation with light microscopy.

We illustrate here how to use electron cryotomography (ECT) to study the ultrastructure of bacterial cells in near-native states, to "macromolecular" (~4 nm) resolution.

While much is already known about the basic metabolism of bacterial cells, many fundamental questions are still surprisingly unanswered, including for ...

Optimizing Sample Preparation for Cryogenic Electron Microscopy

Cryogenic electron microscopy (cryo-EM) has revolutionized structural biology by enabling the study of macromolecular structures in near-native conditions, suspended in vitreous ice. This technique allows for the high-resolution visualization of proteins and other biomolecules without the need for crystallization, offering significant insights into their function and mechanism. Recent advancements in single-particle analysis, coupled with improved computational data processing, have made cryo-EM an indispensable tool in modern structural biology. Despite its growing adoption, cryo-EM faces persistent challenges that can limit its effectiveness, particularly uneven particle distribution. This issue often leads to poor resolution and reduced accuracy in reconstructed protein structures. This article outlines a simple, practical approach to address this challenge, using the small heat-shock protein from Methanocaldococcus jannaschii (MjsHSP16.5) as an example. The method optimizes sample preparation to minimize preferential adsorption, ensuring more homogeneous particle distribution and higher-quality protein cryo-EM structures. This technique offers valuable guidance for researchers aiming to overcome similar challenges in structural studies.

This protocol presents a practical method using Methanocaldococcus jannaschii (MjsHSP16.5) as an example to address uneven particle distribution through sample preparation optimization, providing a reference for researchers to efficiently elucidate macromolecular structures using cryogenic electron microscopy (cryo-EM).

Cryogenic electron microscopy (cryo-EM) has revolutionized structural biology by enabling the study of macromolecular structures in near-native ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...

Cryo-Structured Illumination Microscopic Data Collection from Cryogenically Preserved Cells

Three-dimensional (3D) structured illumination microscopy (SIM) allows imaging of fluorescently labelled cellular structures at higher resolution than conventional fluorescence microscopy. This super-resolution (SR) technique enables visualization of molecular processes in whole cells and has the potential to be used in conjunction with electron microscopy and X-ray tomography to correlate structural and functional information. A SIM microscope for cryogenically preserved samples (cryoSIM) has recently been commissioned at the correlative cryo-imaging beamline B24 at the UK synchrotron. 
It was designed specifically for 3D imaging of biological samples at cryogenic temperatures in a manner compatible with subsequent imaging of the same samples by X-ray microscopy methods such as cryo-soft X-ray tomography. This video article provides detailed methods and protocols for successful imaging using the cryoSIM. In addition to instructions on the operation of the cryoSIM microscope, recommendations have been included regarding the choice of samples, fluorophores, and parameter settings. The protocol is demonstrated in U2OS cell samples whose mitochondria and tubulin have been fluorescently labelled.

This protocol demonstrates how to image biological cryo-preserved samples using cryo-structured illumination microscopy. We demonstrate the methodology by imaging the cytoskeleton of U2OS cells.

Three-dimensional (3D) structured illumination microscopy (SIM) allows imaging of fluorescently labelled cellular structures at higher resolution than ...

Regulating Temperature in the Lab: Preserving Samples Using Cold

Preservation of laboratory samples, specimens, and reagents using extreme cold is routinely performed in biomedical research labs. This video will discuss some of the methods for keeping laboratory samples cold and will explain the correct cooling method to use for each experimental requirement.
For example, cooling agents, such as ice and dry ice, are typically used when keeping samples cold during experiments. This video discusses the physical properties of the most commonly used cooling agents, as well as safety precautions for working with them.
When it comes to keeping samples cold in between experiments, cooling equipment, including laboratory grade refrigerators and freezers can be used to preserve samples for extended period of time. Also discussed in this video are types of samples and reagents that can be stored in the commonly-available laboratory cooling equipment.
Finally, the concept of cryopreservation is introduced as a process through which tissues, cells, and biomolecules are cooled to sub-zero temperatures, thereby effectively stopping all sample-degrading biological activity. Several methods of cryopreservation are discussed that minimize or eliminate the formation of damaging ice crystals.

Preserving Samples Using Cold: Cooling Agents and Instruments, and Cryopreservation

Preservation of laboratory samples, specimens, and reagents using extreme cold is routinely performed in biomedical research labs.  This video will ...

Cryogenic Sample Loading into a Magic Angle Spinning Nuclear Magnetic Resonance Spectrometer that Preserves Cellular Viability

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