Research in the Gunjan lab is supported by funding from the National Institutes of Health, National Science Foundation and the Florida Department of Health. We thank undergraduate student John Parker for technical assistance.

1. Preparation of Yeast Popcorn
<ol>
	<li>Grow yeast cells in 0.5 L of YPD media to a density of 1 x 107 cells/mL. Count cells using a Coulter Counter or any other means.</li>
	<li>Centrifuge cells for 10 min at 2,400 g and 4 &#176;C.</li>
	<li>Wash each sample once with 500 mL of ice-cold deionized 18 mega Ohm Milli-Q water.</li>
	<li>Resuspend pellets in 15 mL of ice-cold Lysis Buffer [20 mM HEPES-KOH pH 7.5, 110 mM KCl, 0.1% Tween, 10% glycerol, with freshly added reducing agent 10 mM &#946;-mercaptoethanol, protease inhibitor cocktail, 10 &#956;M proteasome inhibitor MG-132, 1 mM deacetylase inhibitor sodium butyrate and phosphatase inhibitors (1 mM sodium vanadate, 50 mM sodium fluoride, 50 mM sodium &#946;-glycerophosphate)]. See the&#160;Table of Materials&#160;for additional details. Keep resuspended cells on ice until the samples are ready for the next step. 
	NOTE: A wide variety of lysis buffers containing a number of non-ionic detergents can be used with the samples and specific inhibitors included in it based on the downstream application. Inclusion of protease and proteasomal inhibitors are crucial for preventing protein degradation, especially once the extracts are thawed.</li>
	<li>Make snap frozen yeast popcorn by slowly adding the cell suspension one drop at a time using a pre-chilled serological pipette into one or more 50 mL centrifuge tubes kept on dry ice and filled with LN2 until just below the rim. Top up liquid nitrogen in the tube frequently to compensate for its loss due to rapid evaporation. Work in a well-ventilated area to avoid the hazards associated with nitrogen asphyxiation.
	<ol>
		<li>Instead of a 50 mL tube, use multiple 50 mL tubes tied together (or any container with a large mouth, such as a large plastic centrifuge bottle that can tolerate cryogenic temperatures) for pop-corn preparation (Figure 1A). The larger the container and its opening, the easier is the popcorn preparation as this prevents clumping of the popcorn to form large aggregates. A size range of 0.3-0.5 cm for the popcorn is ideal for achieving proper grinding/pulverization using the freezer mill (Figure 1B).</li>
		<li>Ensure that the LN2 has evaporated completely from the tubes containing the popcorn prior to storing them at -80 &#176;C. It is possible to stop after preparing the yeast popcorn as they are stable for several years when maintained at -80 &#176;C.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61164/61164fig01.jpg" /> 
Figure 1:&#160;Yeast popcorn preparation.&#160;(A)&#160;Yeast &#34;popcorn&#34; is made by the dropwise freezing of the cell suspension in LN2. We use one to three 50 mL tubes held together with a rubber band and placed in an ice bucket filled with dry ice. The tubes are filled with LN2&#160;until just below their rims and are topped up with liquid nitrogen frequently to keep them nearly full until all the cell suspension had been made into popcorn (B)&#160;The size of the popcorn is an important determinant of optimal grinding efficiency. The size range of the popcorn should be between 0.3 and 0.5 cm in diameter.&#160;<a href="https://www.jove.com/files/ftp_upload/61164/61164fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Cryogrinding
<ol>
	<li>To start, ensure that there is enough LN2 available to chill the freezer mill, grinding vials, supplies and to operate the machine for all samples. For less than five samples, 30-35 L of LN2 should suffice. Fill the freezer mill chamber with LN2 up to the fill line. 
	WARNING: All steps involving LN2 should be performed in a well-ventilated area and the freezer mill itself should be located in such an area to avoid risk of asphyxiation. Wear personal protective equipment including proper footwear, lab coat, safety glasses, a base layer of nitrile gloves, then thermal gloves, followed by another pair of nitrile gloves. Use extreme caution when handling LN2.</li>
	<li>Close the freezer mill lid slowly, avoiding the splash of LN2 and allow a few min for the machine to cool down. It may be necessary to refill the chamber with LN2 to compensate for the loss due to evaporation before proceeding to the next step. 
	NOTE: Refill LN2 as needed only up to the fill line. Do not overfill the chamber as this can be both dangerous and detrimental to the machine. Automated refill systems are available for freezer mill that can replenish the evaporating LN2 as needed directly from a storage tank connected to the freezer mill.</li>
	<li>Pre-chill the large grinding vials and the magnetic impactor bar by dunking them in LN2 kept in a separate small dewar. Decant all the LN2 when the LN2 stops bubbling. Then add the sample/yeast popcorn to the grinding vial and seal it tightly with the two stainless steel end plugs.
	<ol>
		<li>Do not fill more than one-third of the grinding vial with the sample as that can reduce the efficiency of grinding. Instead, larger samples can be processed by dividing them into two or more vials and grinding them sequentially. Smaller grinding vials can be used for smaller samples (up to 3 mL).</li>
	</ol>
	</li>
	<li>Place the grinding vial in the freezer mill chamber and lock it in place. Close the lid.</li>
	<li>For budding yeast, grind the samples for a total of three cycles for 2 min/cycle (with 2 min break for cooling between cycles) at a crushing rate of 14. When the machine stops after completing the cycles, open the freezer mill lid slowly and carefully unlock the vial with the powdered frozen cell lysate and remove it from the freezer mill. If multiple small vials are used, work quickly to remove one vial at a time and place them in dry ice. 
	NOTE: There is usually no need to adjust the grinding parameters when using vials of different sizes with the same type of sample. However, the grinding parameters such as the number of cycles and the grinding rate need to be empirically determined for different sample types, depending on the ease with which they lyse. Most mammalian cells and soft tissues will lyse in 1-2 cycles at a crushing rate of 10. Harder to lyse samples such as bacteria, yeast, fly larvae and adult fruit flies require 3-6 cycles at the maximum rate, while hard tissues such as bone, teeth, etc. may require up to 10 cycles. A small aliquot of the thawed sample can be viewed under the microscope before and after grinding to count the number of intact, unlysed cells to determine the lysis efficiency. An excellent analysis of the impact of cryogrinding parameters on the release of proteins and DNA from budding yeast has been published previously15.</li>
	<li>Working quickly so as not to allow the sample to thaw out, carefully unscrew one of the end pieces (pick the one that appears to have become somewhat loose during the grinding) using the opening tool. Then, use a pair of long forceps (that are pre-chilled in a bucket with dry ice) to remove the impactor bar. Collect the pulverized sample by inverting and tapping the grinding vial onto a polystyrene weighing dish pre-chilled with LN2 and kept on dry ice.</li>
	<li>Once all of the powdered frozen cell lysate has been recovered from the grinding vial, pour the powdered lysate from the weighing dish back into the 50 mL tube and proceed immediately to the slow thawing step described next for best results. Alternatively, store the frozen powdered lysate overnight at -80 &#176;C, although this may result in some degradation.</li>
	<li>Prepare an ice bucket with a 50% slurry of ice and water and place it on a stir plate with a magnetic stirrer. Submerge a wire rack or another suitable rack in the ice slurry to hold the samples.
	<ol>
		<li>Then, slowly thaw the samples on an ice slurry bath with constant agitation of the slurry using a magnetic stirrer. Add more ice to replace the melting ice (some water may need to be discarded to prevent the ice bucket from overflowing). Since several inhibitors have short half lives in aqueous buffers, add additional protease inhibitor cocktail (see Table of Materials) and 10 &#956;M proteasome inhibitor MG-132 to the lysate once the samples start thawing (after approximately 30 min).</li>
		<li>Remove ice formed on the outside of the tubes every 5 min to expedite the thawing process. Note that rapid thawing of the samples at room temperature or higher can lead to significant degradation.</li>
	</ol>
	</li>
	<li>After the samples have thawed completely (it may take well over an hour depending on the amount of sample), centrifuge the lysate at ~3,220 g for 20 min in a refrigerated tabletop centrifuge at 4 &#176;C to remove the bulk of the cell debris from the lysate.</li>
	<li>Transfer the supernatant to 50 mL polycarbonate centrifuge tubes that have been pre-chilled on ice and centrifuge the samples at 16,000 g for 20 min at 4 &#176;C (see Table of Materials).</li>
	<li>Transfer only the clear supernatant consisting of the clarified whole cell extract from the center of the liquid column in the tube very slowly and carefully, without disturbing the cloudy lipid layer (Figure 2), into a chilled 15 mL tube using pre-chilled serological pipettes. Avoid removing all of the lysate in the centrifuge tube to prevent carryover of lipids and debris. The leftover lysate in the centrifuge tube can be centrifuged multiple times to recover small amounts of extract after each spin. 
	&#8203;NOTE: Lipids along with any denatured proteins would appear cloudy/milky and are located at the top of the liquid/air interface, and occasionally above the pellet itself that consists of insoluble cellular debris (Figure 2).</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61164/61164fig02.jpg" /> 
Figure 2:&#160;Extract layers in a centrifuge tube.&#160;The major visible features of the whole cell extract in a tube following centrifugation at 16,000 g for 20 min are indicated. The relative abundance of each feature depends on the sample type, the growth phase of the cells (exponential versus stationary), the amount of lysis buffer used to resuspend cells and the lysis efficiency.&#160;<a href="https://www.jove.com/files/ftp_upload/61164/61164fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="12">
	<li>Centrifuge any remaining lysate for 5 min to recover more of the extract. Repeat this 5 min centrifugation multiple times if needed to recover as much of the clear extract as possible. Discard the cloudy lysate containing lipids close to the bottom of the tube near the pellet.</li>
	<li>Use these extracts for downstream applications such as protein complex purification, immunoprecipitation and proteomic analysis.</li>
</ol>

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The authors have nothing to disclose.

A limitation of studying native proteins from yeast is the inefficient lysis of yeast cells due to&#160;their tough cell wall. Although several methods have been developed, the most consistent and efficient method in our hands is the cryogrinding of yeast cells flash frozen as popcorn. This method allows the reliable preparation of high-quality whole cell extracts from budding yeast compared to other lysis methods. The representative results demonstrated that cryogrinding is superior to a popular mechanical method for ye...

Yeast is a popular model organism for protein studies, as it is a simple eukaryotic organism with an abundance of genetic and biochemical tools available for researchers1. Because of their sturdy cell wall, one challenge that researchers face is in efficiently lysing the cells without damaging the cellular contents. Different methods are available for obtaining protein extracts through disruption of yeast cells which include enzymatic lysis (zymolyase)2,3, chemical lysis4, physical lysis by freeze-thaw5, pressure-based (French press)6,7, mechanical (glass beads, coffee grinder)8,9, sonication-based10 and cryogenic2,11. The efficiency of cell lysis and the protein yield can vary considerably depending on the technique employed, thus affecting the end result or suitability for the desired downstream application for the lysate. When studying proteins that are unstable, have fleeting posttranslational modifications, or are temperature sensitive, it is particularly important to use a method that will minimize sample loss or degradation during preparation.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Extract preparation technique</td>
			<td>Details</td>
			<td>Advantages</td>
			<td>Disadvantages</td>
			<td>Downstream analysis</td>
			<td>Reference</td>
		</tr>
		<tr>
			<td>French press: High-pressure homogenizer (aka Microfluidizer) with enzymatic pretreatment using Zymolyase</td>
			<td>Zymolyase-20T, a Microfluidizer high-pressure homogenizer. The disruptor consists of an air-driven, high-pressure pump (ratio 1:250; required air pressure 0.6-l MPa) and a special disruption chamber with an additional back pressure unit. A minimum sample size of 20 mL is required for processing.</td>
			<td>Final total disruption obtained using the combined protocol approached 100 % with 4 passes at a pressure of 95 MPa, as compared to only 32 % disruption with 4 passes at 95 MPa using only homogenization without the Zymolyase.</td>
			<td>Not appropriate for small scale applications. The enzymes can get expensive for large scale preparations.</td>
			<td>Protein purification</td>
			<td>6</td>
		</tr>
		<tr>
			<td>Bead beater: Zymolyase treated cells lysed with glass beads in a fastprep instrument</td>
			<td>Roughly an equal volume of cold, dry, acid-washed 0.5 mm glass beads is added to a given volume of cell pellet in lysis buffer and the cells are disrupted by vigorous manual agitation.</td>
			<td>It is particularly useful when making extracts from many different small yeast cultures for assaying purposes rather than for protein purification.</td>
			<td>During the glass bead procedure, proteins are treated harshly causing extensive foaming leading to protein denaturation. The amount of cell breakage varies, while proteolysis as well as modification of the proteins may result from heating of the extract above 4&#176;C during the mechanical breakage.</td>
			<td>Mostly DNA &#38; RNA analyses, but also protein analysis by denaturing gel electropheoresis, either with or without Western blotting.</td>
			<td>8</td>
		</tr>
		<tr>
			<td>Zymolyase treatment&#160;&#160; followed by lysis using a combination of osmotic shock and Dounce homogenization</td>
			<td>After enzymatic digestion of cell walls, spheroplasts are lysed with 15 to 20 strokes of a tight-fitting pestle (clearance 1 to 3 &#181;m) in a Dounce homogenizer.</td>
			<td>Advantageous to use protease-deficient strains such as BJ926 or EJ101. This is the gentlest way to break yeast cells and hence it is most suitable for preparing extracts that can carry out complex enzymatic functions (e.g., translation, transcription, DNA replication) and in which the integrity of macromolecular structures (e.g., ribosomes, splicesomes) has to be maintained. It is also useful for isolating intact nuclei that can be used for chromatin studies (Bloom and Carbon, 1982) or for nuclear protein extracts (Lue and Kornberg, 1987).</td>
			<td>The major disadvantages of the spheroplast lysis procedure are that it is relatively tedious and expensive, especially for large-scale preparations (&#62;10 liters), and the long incubation periods can lead to proteolysis or protein modification. &#160;For chromatin preparations, they seem to be of varying or lower quality than those produced by the differential centrifugation (based on nucleosome ladder integrity).</td>
			<td>Isolating intact nuclei for chromatin studies, extracts that can carry out complex enzymatic functions, extracts requiring the integrity of 
			macromolecular structures, nuclear protein extracts.</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Cell Disruption of flash frozen cells by grinding in Liquid Nitrogen using a mortar/pestle or a blender</td>
			<td>Cells are frozen immediately in liquid nitrogen and then lysed by grinding manually in a mortar using a pestle, or using a Waring blender in the presence of liquid nitrogen.</td>
			<td>The protocol is quick and easy. It can accommodate varying amounts of yeast cells including very large cultures. Its main advantage is that cells are taken immediately from the actively growing state into liquid nitrogen (&#8722;196&#176;C), decreasing degradative enzyme activities such as proteases and nucleases as well as activities that modify proteins (e.g., phosphatases and kinases). It is particularly suited for making whole-cell extracts from a single yeast culture for large-scale protein purification.&#160;</td>
			<td>A bit messy and potentially dangerous to the careless investigator. Small samples (i.e., 10- to 100-ml yeast cultures) are not easily processed because there is not enough mass of frozen cell clumps to fracture effectively in the blender. It is time-consuming to process individual samples and to clean the equipment between uses.</td>
			<td>Whole-cell extracts from a single yeast culture for large-scale protein purification.</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Autolysis, Bead mill</td>
			<td>pH 5.0, 50 &#176;C, 24 h, 200 rpm / &#216; 0.5 mm, 5 &#215; 3 min/3 min</td>
			<td>Quick and efficient lysis, especially for small scale extract preparation</td>
			<td>Heat generation leads to denaturation and degradation of macromolecules. Bead beating equipment required.</td>
			<td>Small scale analyses.</td>
			<td rowspan="2">10</td>
		</tr>
		<tr>
			<td>Autolysis, Sonication</td>
			<td>pH 5.0, 50 &#176;C, 24 h, 200 rpm, 4 &#215; 5 min/2 min, pulser 80%, power 80%</td>
			<td>Sonication equipment is usually available in most institutions.</td>
			<td>Heat generation leads to denaturation and degradation of macromolecules. Sonication equipment required. Slow lysis can take more than 24 hours.</td>
			<td>Yeast cell wall preparations.</td>
		</tr>
		<tr>
			<td>Boiling and freeze-thaw process</td>
			<td>No specialized equipment needed other than a standard freezer and a heating block or hot water bath.</td>
			<td>Efficient, reproducible, simple and inexpensive.</td>
			<td>Heat generation leads to denaturation and degradation of macromolecules.</td>
			<td>DNA analyses by PCR.</td>
			<td>5</td>
		</tr>
	</tbody>
</table>
Table 1:&#160;Comparison of methods available for the preparation of yeast extracts.
Cryogrinding (aka cryogenic grinding/cryogenic milling) is commonly employed to retrieve nucleic acids, proteins or chemicals from temperature sensitive samples in a reliable manner for quantitative or qualitative analyses. It has been used successfully for multiple applications in diverse fields including biotechnology, toxicology, forensic science12,13, environmental science, plant biology14 and food science. Isolation of intact biological macromolecules is usually critically dependent on the temperature. Extremely low temperatures ensure that the proteases and nucleases stay inactive, resulting in a reliable isolation of intact proteins, nucleic acids and other macromolecules for subsequent analyses. Indeed, a freezer mill typically maintains a sample temperature of -196 &#176;C (the boiling point of LN2), thus minimizing DNA/RNA or protein denaturation and degradation.
The freezer mill employs an electromagnetic grinding chamber that rapidly moves a solid metal bar or cylinder back and forth within a vial containing the sample to be pulverized between stainless steel end plugs. The instrument creates and rapidly reverses a magnetic field within the grinding chamber. As the magnetic field shifts back and forth, the magnet crushes the sample against the plugs thus achieving the &#39;cryogrinding&#39; and the pulverization of the popcorn. The freezer mill replaces the mortar and pestle and allows the sequential processing of multiple samples (or up to 4 smaller samples simultaneously) with high reproducibility and avoids the user-to-user variability associated with manual grinding. Once the samples are processed, the cell extracts can be used for a variety of downstream applications.

<table><tbody><tr><td>50 mL polycarbonate tubes with screw caps</td><td>Beckman</td><td>357002</td><td>Centrifuge tubes</td></tr><tr><td>BD Bacto Peptone</td><td>BD Biosiences</td><td>211677</td><td>Yeast YPD media component</td></tr><tr><td>BD Bacto Yeast Extract</td><td>BD Biosiences</td><td>212750</td><td>Yeast YPD media component</td></tr><tr><td>Beckman Avanti centrifuge</td><td>Beckman</td><td>B38624</td><td>High speed centrifuge</td></tr><tr><td>Beckman JLA-9.1000</td><td>Beckman</td><td>366754</td><td>Rotor</td></tr><tr><td>D-(+)-Dextrose Anhydrous</td><td>MP Biomedicals</td><td>901521</td><td>Yeast YPD media component</td></tr><tr><td>Eppendorf A-4-44</td><td>Eppendorf</td><td>22637461</td><td>Swinging bucket rotor</td></tr><tr><td>Eppendorf refrigerated centrifuge 5810 R</td><td>Eppendorf</td><td>22625101</td><td>Refrigerated centrifuge</td></tr><tr><td>Glycerol</td><td>SIGMA-ALDRICH</td><td>G5150-1GA</td><td>Volume excluder and cryoprotectant</td></tr><tr><td>HEPES</td><td>FisherBiotech</td><td>BP310-100</td><td>Buffer</td></tr><tr><td>HIS6 antibody</td><td>Novagen</td><td>70796</td><td>Antibody for HIS tag</td></tr><tr><td>KCl</td><td>SIGMA-ALDRICH</td><td>P9541-1KG</td><td>Salt for maintaining ionic strength</td></tr><tr><td>MG-132</td><td>CALBIOCHEM</td><td>474790</td><td>Proteasome Inhibitor</td></tr><tr><td>Phosphatase inhibitor cocktail</td><td>ThermoFisher Scientific</td><td>A32957</td><td>Phosphatase inhibitor cocktail</td></tr><tr><td>Ponceau S</td><td>SIGMA</td><td>P7170-1L</td><td>Protein Stain</td></tr><tr><td>Protease inhibitor cocktail</td><td>ThermoFisher Scientific</td><td>A32963</td><td>Protease inhibitor cocktail</td></tr><tr><td>Rotor JLA 25.500</td><td>Beckman</td><td>JLA 25.500</td><td>Rotor</td></tr><tr><td>Sodium Butyrate</td><td>EM Science</td><td>BX2165-1</td><td>Histone Deacetylase Inhibitor</td></tr><tr><td>Sodium Fluoride</td><td>Sigma-Aldrich</td><td>S6521</td><td>Phosphatase Inhibitor</td></tr><tr><td>Sodium Vanadate</td><td>MP Biomedicals</td><td>159664</td><td>Phosphatase Inhibitor</td></tr><tr><td>Sodium &amp;beta;-glycerophosphate</td><td>Alfa Aesar</td><td>13408-09-8</td><td>Phosphatase Inhibitor</td></tr><tr><td>Spex Certiprep 6850 freezer mill</td><td>SPEX Sample Prep</td><td>6850</td><td>Freezer Mill</td></tr><tr><td>TALON Metal Affinity Resin</td><td>BD Biosiences</td><td>635502</td><td>For pulling down HIS tagged proteins</td></tr><tr><td>Tween 20</td><td>VWR International</td><td>VW1521-07</td><td>Non-ionic detergent</td></tr><tr><td>&amp;beta;-Mercaptoethanol</td><td>AMRESCO</td><td>M131-250ML</td><td>Reducing agent</td></tr></tbody></table>

preparation of cell extracts by cryogrinding in an automated freezer mill

The ease of genetic manipulation and the strong evolutionary conservation of eukaryotic cellular machinery in the budding yeast Saccharomyces cerevisiae has made it a pre-eminent genetic model organism. However, since efficient protein isolation depends upon optimal disruption of cells, the use of yeast for biochemical analysis of cellular proteins is hampered by its cell wall which is expensive to digest enzymatically (using lyticase or zymolyase), and difficult to disrupt mechanically (using a traditional bead beater, a French press or a coffee grinder) without causing heating of samples, which in turn causes protein denaturation and degradation. Although manual grinding of yeast cells under liquid nitrogen (LN2) using a mortar and pestle avoids overheating of samples, it is labor intensive and subject to variability in cell lysis between operators. For many years, we have been successfully preparing high quality yeast extracts using cryogrinding of cells in an automated freezer mill. The temperature of -196 &#176;C achieved with the use of LN2 protects the biological material from degradation by proteases and nucleases, allowing the retrieval of intact proteins, nucleic acids and other macromolecules. Here we describe this technique in detail for budding yeast cells which involves first freezing a suspension of cells in a lysis buffer through its dropwise addition into LN2 to generate frozen droplets of cells known as "popcorn". This popcorn is then pulverized under LN2 in a freezer mill to generate a frozen "powdered" extract which is thawed slowly and clarified by centrifugation to remove insoluble debris. The resulting extracts are ready for downstream applications, such as protein or nucleic acid purification, proteomic analyses, or co-immunoprecipitation studies. This technique is widely applicable for cell extract preparation from a variety of microorganisms, plant and animal tissues, marine specimens including corals, as well as isolating DNA/RNA from forensic and permafrost fossil specimens.

We compared two different methods for yeast cell lysis, namely glass bead milling at 4 &#176;C and an automated cryogrinding method at -196 &#176;C, to assess the relative recovery proteins in the cell extracts prepared with both methods. For this study, we chose to use a budding yeast strain YAG 1177 (MAT a lys2-810 leu2-3,-112 ura3-52 his3-&#916;200 trp1-1[am] ubi1-&#916;1::TRP1 ubi2-&#916;2::ura3 ubi3-&#916;ub-2 ubi4-&#916;2::LEU2 [pUB39] [pUB221])16 carrying a ...

Watch this Scientific Journal Video about Preparation of Cell Extracts by Cryogrinding in an Automated Freezer Mill at JoVE.com

Preparation of Cell Extracts by Cryogrinding in an Automated Freezer Mill

We describe a reliable method for the preparation of whole cell extracts from yeast or other cells using a cryogenic freezer mill that minimizes degradation and denaturation of proteins. The cell extracts are suitable for purification of functional protein complexes, proteomic analyses, co-immunoprecipitation studies and detection of labile protein modifications.

The ease of genetic manipulation and the strong evolutionary conservation of eukaryotic cellular machinery in the budding yeast Saccharomyces cerevisiae ...

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Research

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Biology

8.0K Views.  Florida State University College of Medicine. We describe a reliable method for the preparation of whole cell extracts from yeast or other cells using a cryogenic freezer mill that minimizes degradation and denaturation of proteins. The cell extracts are suitable for purification of functional protein complexes, proteomic analyses, co-immunoprecipitation studies and detection of labile protein modifications.

Video: Preparation of Cell Extracts by Cryogrinding in an Automated Freezer Mill

Protein Complex Affinity Capture from Cryomilled Mammalian Cells

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this technique is also frequently referred to as immunoprecipitation. Affinity capture can be applied in a bench-scale and in a high-throughput context. When coupled with protein mass spectrometry, affinity capture has proven to be a workhorse of interactome analysis. Although there are potentially many ways to execute the numerous steps involved, the following protocols implement our favored methods. Two features are distinctive: the use of cryomilled cell powder to produce cell extracts, and antibody-coupled paramagnetic beads as the affinity medium. In many cases, we have obtained superior results to those obtained with more conventional affinity capture practices. Cryomilling avoids numerous problems associated with other forms of cell breakage. It provides efficient breakage of the material, while avoiding denaturation issues associated with heating or foaming. It retains the native protein concentration up to the point of extraction, mitigating macromolecular dissociation. It reduces the time extracted proteins spend in solution, limiting deleterious enzymatic activities, and it may reduce the non-specific adsorption of proteins by the affinity medium. Micron-scale magnetic affinity media have become more commonplace over the last several years, increasingly replacing the traditional agarose- and Sepharose-based media. Primary benefits of magnetic media include typically lower non-specific protein adsorption; no size exclusion limit because protein complex binding occurs on the bead surface rather than within pores; and ease of manipulation and handling using magnets.

Here we describe protocols to disrupt mammalian cells by solid-state milling at a cryogenic temperature, produce a cell extract from the resulting cell powder, and isolate protein complexes of interest by affinity capture upon antibody-coupled micron-scale paramagnetic beads.

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this ...

Biochemistry

Optimization and Comparative Analysis of Plant Organellar DNA Enrichment Methods Suitable for Next-generation Sequencing

Plant organellar genomes contain large, repetitive elements that may undergo pairing or recombination to form complex structures and/or sub-genomic fragments. Organellar genomes also exist in admixtures within a given cell or tissue type (heteroplasmy), and an abundance of subtypes may change throughout development or when under stress (sub-stoichiometric shifting). Next-generation sequencing (NGS) technologies are required to obtain deeper understanding of organellar genome structure and function. Traditional sequencing studies use several methods to obtain organellar DNA: (1) If a large amount of starting tissue is used, it is homogenized and subjected to differential centrifugation and/or gradient purification. (2) If a smaller amount of tissue is used (i.e., if seeds, material, or space is limited), the same process is performed as in (1), followed by whole-genome amplification to obtain sufficient DNA. (3) Bioinformatics analysis can be used to sequence the total genomic DNA and to parse out organellar reads. All these methods have inherent challenges and tradeoffs. In (1), it may be difficult to obtain such a large amount of starting tissue; in (2), whole-genome amplification could introduce a sequencing bias; and in (3), homology between nuclear and organellar genomes could interfere with assembly and analysis. In plants with large nuclear genomes, it is advantageous to enrich for organellar DNA to reduce sequencing costs and sequence complexity for bioinformatics analyses. Here, we compare a traditional differential centrifugation method with a fourth method, an adapted CpG-methyl pulldown approach, to separate the total genomic DNA into nuclear and organellar fractions. Both methods yield sufficient DNA for NGS, DNA that is highly enriched for organellar sequences, albeit at different ratios in mitochondria and chloroplasts. We present the optimization of these methods for wheat leaf tissue and discuss major advantages and disadvantages of each approach in the context of sample input, protocol ease, and downstream application.

The comparison and optimization of two plant organellar DNA enrichment methods are presented: traditional differential centrifugation and fractionation of the total gDNA based on methylation status. We assess the resulting DNA quantity and quality, demonstrate performance in short-read next-generation sequencing, and discuss the potential for use in long-read single-molecule sequencing.

Plant organellar genomes contain large, repetitive elements that may undergo pairing or recombination to form complex structures and/or sub-genomic ...

Genetics

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 ...

Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could easily be adapted for other biological samples. The basic principles for preparing samples, milling, and viewing lamellae are common to all instruments and the protocol can be followed as a general guide to on-grid cryo-lamella preparation for cryo-electron microscopy (cryoEM) and cryo-electron tomography (cryoET). Electron microscopy grids supporting the cells are plunge-frozen into liquid nitrogen-cooled liquid ethane using a manual or automated plunge freezer, then screened on a light microscope equipped with a cryo-stage. Frozen grids are transferred into a cryo-scanning electron microscope equipped with a focused ion beam (cryoFIB-SEM). Grids are routinely sputter coated prior to milling, which aids dispersal of charge build-up during milling. Alternatively, an e-beam rotary coater can be used to apply a layer of carbon-platinum to the grids, the exact thickness of which can be more precisely controlled. Once inside the cryoFIB-SEM an additional coating of an organoplatinum compound is applied to the surface of the grid via a gas injection system (GIS). This layer protects the front edge of the lamella as it is milled, the integrity of which is critical for achieving uniformly thin lamellae. Regions of interest are identified via SEM and milling is carried out in a step-wise fashion, reducing the current of the ion beam as the lamella reaches electron transparency, in order to avoid excessive heat generation. A grid with multiple lamellae is then transferred to a transmission electron microscope (TEM) under cryogenic conditions for tilt-series acquisition. A robust and contamination-free workflow for lamella preparation is an essential step for downstream techniques, including cellular cryoEM, cryoET, and sub-tomogram averaging. Development of these techniques, especially for lift-out and milling of high-pressure frozen samples, is of high-priority in the field.

Using focused ion beam milling to produce vitreous on-grid lamellae from plunge frozen biological samples for cryo-electron tomography.

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could ...

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 ...

Chemistry

Manual Blot-and-Plunge Freezing of Biological Specimens for Single-Particle Cryogenic Electron Microscopy

Imaging biological specimens with electrons for high-resolution structure determination by single-particle cryogenic electron microscopy (cryoEM) requires a thin layer of vitreous ice containing the biomolecules of interest. Despite numerous technological advances in recent years that have propelled single-particle cryoEM to the forefront of structural biology, the methods by which specimens are vitrified for high-resolution imaging often remain the rate-limiting step. Although numerous recent efforts have provided means to overcome hurdles frequently encountered during specimen vitrification, including the development of novel sample supports and innovative vitrification instrumentation, the traditional manually operated plunger remains a staple in the cryoEM community due to the low cost to purchase and ease of operation. Here, we provide detailed methods for using a standard, guillotine-style manually operated blot-and-plunge device for the vitrification of biological specimens for high-resolution imaging by single-particle cryoEM. Additionally, commonly encountered issues and troubleshooting recommendations for when a standard preparation fails to yield a suitable specimen are also described.

This manuscript outlines the blot-and-plunge method to manually freeze biological specimens for single-particle cryogenic electron microscopy.

Imaging biological specimens with electrons for high-resolution structure determination by single-particle cryogenic electron microscopy (cryoEM) requires ...

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 ...

Extracting Modified Microtubules from Mammalian Cells to Study Microtubule-Protein Complexes by Cryo-Electron Microscopy

Microtubules are an important part of the cytoskeleton and are involved in intracellular organization, cell division, and migration. Depending on the posttranslational modifications, microtubules can form complexes with various interacting proteins. These microtubule-protein complexes are often implicated in human diseases. Understanding the structure of such complexes is useful for elucidating their mechanisms of action and can be studied by cryo-electron microscopy (cryo-EM). To obtain such complexes for structural studies, it is important to extract microtubules containing or lacking specific posttranslational modifications. Here, we describe a simplified protocol to extract endogenous tubulin from genetically modified mammalian cells, involving microtubule polymerization, followed by sedimentation using ultracentrifugation. The extracted tubulin can then be used to prepare cryo-electron microscope grids with microtubules that are bound to a purified microtubule-binding protein of interest. As an example, we demonstrate the extraction of fully tyrosinated microtubules from cell lines engineered to lack the three known tubulin-detyrosinating enzymes. These microtubules are then used to make a protein complex with enzymatically inactive microtubule-associated tubulin detyrosinase on cryo-EM grids.

Here, we describe a protocol to extract endogenous tubulin from mammalian cells, which can lack or contain specific microtubule-modifying enzymes, to obtain microtubules enriched for a specific modification. We then describe how the extracted microtubules can be decorated with purified microtubule-binding proteins to prepare grids for cryo-electron microscopy.

Microtubules are an important part of the cytoskeleton and are involved in intracellular organization, cell division, and migration. Depending on the ...

Preparation of Electron Microscopy Grids for In Situ Cellular Cryotomography

Sample Preparation for In Situ Cryotomography of Mammalian Cells

In situ cellular cryotomography is a powerful technique for studying complex objects in their native frozen-hydrated cellular context, making it highly relevant to cellular biology and virology. The potential of combining cryotomography with other microscopy modalities makes it a perfect technique for integrative and correlative imaging. However, sample preparation for in situ cellular tomography is not straightforward, as cells do not readily attach and stretch over the electron microscopy grid. Additionally, the grids themselves are fragile and can break if handled too forcefully, resulting in the loss of imageable areas. The geometry of tissue culture dishes can also pose a challenge when manipulating the grids with tweezers. Here, we describe the tips and tricks to overcome these (and other) challenges and prepare good-quality samples for in situ cellular cryotomography and correlative imaging of adherent mammalian cells. With continued advances in cryomicroscopy technology, this technique holds enormous promise for advancing our understanding of complex biological systems.

Author Spotlight: Optimizing Grid Preparation for Enhanced Cryoelectron Tomography 

This method provides an accessible and flexible protocol for the preparation of electron microscopy (EM) grids for in situ cellular cryotomography and correlative light and electron microscopy (CLEM).

In situ cellular cryotomography is a powerful technique for studying complex objects in their native frozen-hydrated cellular context, making it highly ...

Sample Preparation by 3D-Correlative Focused Ion Beam Milling for High-Resolution Cryo-Electron Tomography

Cryo-electron tomography (cryo-ET) has become the method of choice for investigating cellular ultrastructure and molecular complexes in their native, frozen-hydrated state. However, cryo-ET requires that samples are thin enough to not scatter or block the incident electron beam. For thick cellular samples, this can be achieved by cryo-focused ion beam (FIB) milling. This protocol describes how to target specific cellular sites during FIB milling using a 3D-correlative approach, which combines three-dimensional fluorescence microscopy data with information from the FIB-scanning electron microscope. Using this technique, rare cellular events and structures can be targeted with high accuracy and visualized at molecular resolution using cryo-transmission electron microscopy (cryo-TEM).

Here, we present a pipeline for 3D-correlative focused ion beam milling on guiding the preparation of cellular samples for cryo-electron tomography. The 3D position of fluorescently tagged proteins of interest is first determined by cryo-fluorescence microscopy, and then targeted for milling. The protocol is suitable for mammalian, yeast, and bacterial cells.

Cryo-electron tomography (cryo-ET) has become the method of choice for investigating cellular ultrastructure and molecular complexes in their native, ...