High-pressure Freezing Followed by Freeze-substitution, an Optimal Electron Microscope Technique to Study Golgi Apparatus Organization and Membrane Trafficking

Abstract

A major goal of structural biologists is to preserve samples as close to their living state as possible. High-pressure freezing (HPF) is a state-of-art technique that freezes the samples at high pressure (~2100 bar) and low temperature (−196°C) within milliseconds. This ultrarapid fixation enables simultaneous immobilization of all cellular components and preserve the samples in a near-native state. This facilitates the study of dynamic processes in Golgi apparatus organization and membrane trafficking. The work in our laboratory shows that high-pressure freezing followed by freeze-substitution (FS), the introduction of organic solvents at low temperature prior to plastic embedding, can better preserve the structure of Golgi apparatus and Golgi-associated vesicles. Here, we present a protocol for freezing monolayer cell cultures on sapphire discs followed by freeze-substitution. We were able to use this protocol to successfully study Golgi organization and membrane trafficking in HeLa cells. The protocol gives decidedly better preservation of Golgi apparatus and associated vesicles than conventional chemically fixed preparation and as a plastic embedded preparation can be readily extended to 3D electron microscopy imaging through sequential block face scanning electron microscopy. The 3D imaging of a multi-micron thick organelle such as the Golgi apparatus located near the cell nucleus is greatly facilitated relative to hydrated sample imaging techniques such as cryo-electron microscopy

1. Introduction

Many biological structures within the Golgi apparatus and associated vesicle trafficking steps are below the 200-nm resolution of light microscopy. Electron microscopy presents a powerful alternative to accurately visualize cellular details at higher resolution. In the traditional processing method, cells are chemically fixed, dehydrated and embedded into a plastic resin [1]. However, this method can introduce significant morphological artifacts. The chemical fixation procedure is relatively slow and not suitable to stop dynamic processes [2,3]. Chemical reagents like glutaraldehyde and organic solvents can influence and change cellular structures and molecules [4,5]. Therefore, high-pressure freezing (HPF) and freeze-substitution (FS) protocols have been developed to overcome these drawbacks and preserve cell ultrastructure close to the normal living state [6].

In the HPF technique, specimens are frozen at high pressure (~2100 bar) in liquid nitrogen (−196°C) within milliseconds. These conditions do not allow water inside the cells to form ice crystals that can damage cell structures. Instead, the water becomes a very viscous fluid [7,8]. The ultrarapid fixation enables simultaneous immobilization of all cellular components and facilitates the study of dynamic processes [9]. HPF is followed by FS, during which water in the sample is replaced by fixatives, dehydrating agents (organic solvents), and stains at low temperature [10]. The organic solvents permeabilize the cells in a near-native state and provide access for later plastic embedding steps. The samples are then brought to room temperature and embedded in plastic resin and further processed for imaging [10]. The plastic embedding facilitates such 3D procedures as serial block face scanning electron microscopy in which depths of 10s of microns can be readily visualized.

Since the first HPF machine (the Balzer’s HPM 010) became commercially available, more and more scientists started to use this technology. Today, HPF becomes a state-of-art technique and has made great contributions to generating reliable data for numerous scientific questions. Currently, there are four different HPF machines available: EM PACT2 and HPM 100 (Leica Microsystems), HPM 010 and HPF Compact 02 (Engineering Office M) [6]. The choice of which to use depends on the kind of sample and on the sample preparation techniques. Here, we present a step-by-step protocol for using a Leica EM PACT2 to freeze monolayer cells. In the EM PACT2 system, specimen carriers are used to hold anything from cells to small tissues. Sapphire discs seeded with monolayer cell cultures fit into the carriers. We were able to successfully freeze monolayer HeLa cell cultures on sapphire discs in EM PACT2 HPF device followed by freeze-substitution in an automated freeze substitution (AFS, Leica Microsystems) system [11–15]. In contrast to chemical fixation, HPF improves preservation of the morphological features of the Golgi apparatus and preserves vesicle arrangement in HeLa cells [11]. This allows us to better understand Golgi apparatus organization and membrane trafficking processes (e.g., Figure 1). HPF can be used to prepare samples for electron tomography and serial block face–scanning electron microscopy (SBF-SEM) to give cell ultrastructure in three-dimensional models [9, 11]. Examples contrasting chemical fixation and HPF/FS protocol are shown in Figure 1.

Figure 1. The ultrastructure of the Golgi apparatus in Rab6 depleted HeLa cells prepared by chemical fixation (A) and by high pressure freezing/freeze substitution (HPF/FS, B).

In both cases, HeLa cells were incubated with siRab6 [11,13] for 4 days to knockdown Rab6 protein. In contrast to chemically fixed cells, HPF/FS gave a long, continuous Golgi cisternae with dilated structures restricted to one side of the Golgi stack. Importantly, apparent vesicles stand out much more prominently in the HPF/FS preparation. We submit that HPF/FS is essential to preserve the native structure of Golgi apparatus in cells.

For illustrative purposes only, we outlined here a generalizable protocol that are specific to Leica Microsystem devices. Exact protocol details will vary from system to system.

• Step 1. Grow cells on sapphire discs

• Step 2. Rinse disc in cryoprotectant at 37°C and transfer to specimen carrier

• Step 3. High pressure freeze cells to quickly preserve structure

• Step 4. Freeze-substitute frozen cells to chemically fix them, introduce organic solvent pre-embedding, and increase membrane contrast

• Step 5. Infiltrate and embed the cells with resin

• Step 6. Analyze samples by transmission electron microscopy (2D imaging), electron tomography or serial block face–scanning electron microscopy (SBF-SEM)

The detailed step-by-step protocol is presented below.

2. Materials

2.1. High-pressure freezing

1. EM PACT2 high-pressure freezer with Rapid Transfer System (RTS) (Leica Microsystems)

2. Essential accessories for EM PACT2/RTS: RTS bajonett loading device, bayonet pod, rapid loader, cryotransfer tool, torque wrench and forceps

3. Carbon coater with which to coat sapphire discs prior to cell culture (Leica Microsystems)

4. Sapphire discs (Leica Microsystems), 1.4 mm in diameter

5. Finder grids (Leica Microsystems), 1.48 mm in diameter

6. Specimen carrier (Leica Microsystems), gold plated, 1.51 mm in diameter, 140 μm deep

7. Stereo microscope with a warmed stage (37°C)

8. Cell culture dishes, culture media and cell incubator

9. DPBS, Dulbecco’s phosphate buffered saline, i.e., calcium and magnesium containing PBS

10. Water bath (37°C)

11. Liquid nitrogen (See Note 1)

2.2. Freeze-substitution

1. Automated freeze-substitution unit (AFS) (Leica Microsystems)

2. Cryovials (2 ml)

3. 15 ml screw cap glass bottles (flat bottom)

4. 12.5% osmium tetroxide (Electron Microscopy Sciences) (See Note 2)

5. 10% glutaraldehyde (Electron Microscopy Sciences) (See Note 2)

6. 10% uranyl acetate stock solution: dissolve 1 g of uranyl acetate in 10 ml of methanol. Store at 4°C in dark (see Notes 2, 3 and 4)

7. Anhydrous acetone (Electron Microscopy Sciences)

8. Molecular grade water

9. Liquid nitrogen

2.3. Infiltration, Embedding and Sectioning

1. Preparation of embedding medium

   1. Fully liquefy Embed-812, Araldite 502 and DDSA by warming these components to 60°C (see Note 2)
   2. Mix 25 ml of Embed-812, 15 ml of Araldite 502 and 55 ml of DDSA in a glass bottle. Place a stir bar and gently stir until the components are completely mixed (see Note 5). Thorough mixing is imperative to be able to obtain uniform resin blocks. The Embed/Araldite mixture can be stored for up to 6 months at 4°C. But freshly prepared mixture is highly recommended.
   3. Immediately before using, warm the Embed/Araldite mixture thoroughly at 60°C. Add 200 μl DMP-30 accelerator for every 10 ml mixture and gently mix it by inverting the tube (see Note 6). DMP-30 turns the mixture to a deep orange color.

2. Microwave processor

3. Flat embedding mold

4. 60°C oven

5. UltraCut-UCT microtome (Leica Microsystems)

6. Single-edge razor blade

7. Diamond knife

8. Grids and grid storage box

9. Chloroform

2.4. Post Staining and Imaging

1. Reynolds lead citrate: to remove any CO2, the molecular grade water should be boiled for 10–15 min prior to use and kept airtight (see Note 7). Add 1.33 g of lead nitrate and 1.76 g of sodium citrate to 30 ml of CO2 free water in a 50 ml volumetric flask. Shake vigorously for 1–2 min and intermittently over 30 min to ensure complete conversion of lead nitrate to lead citrate. Add 8 ml of 1 N NaOH and bring to 50 ml (see Note 8). Invert slowly and store in sealed bottle at 4°C (see Note 9).

2. 0.5% uranyl acetate: dissolve 2.9 g of maleic acid in 400 ml of molecular grade water and adjust pH to 6.0 with 1 N NaOH. In a plastic bottle, dissolve 0.5 g of uranyl acetate dihydrate in 80 ml of maleate buffer and adjust pH to 5.0 with NaOH. Make up to 100 ml with molecular grade water. Store at 4°C in dark (see Notes 2, 3 and 4)

3. Tecnai F20 transmission electron microscope (FEI Co) or other suitable electron microscope operated at 80 keV (for thin sections (50 nm) imaging or at 300 keV (for semi-thick sections (300 nm) imaging and electron tomography.

3. Methods

3.1. Cell Monolayer Culture

1. The 1.4-mm sapphire discs are coated with 10 nm thick carbon layer. Sterilize the coated discs by tissue culture hood ultraviolent radiation for 20–30 min in a 35 mm tissue culture dish (without lid) before using. If the procedure requires selecting a particular cell area, then the sapphire disc can be carbon coated with a finder grid on top to create a reference pattern. The carbon coating procedure facilitates cell growth, separation of the sapphire disc from embedding plastic, and cell re-localization after resin embedding.

2. Add 2 ml cell media in each dish. The discs will float, so gently push them down with sterile forceps (see Notes 10 and 11). The coated sides of discs should always be facing up. Plate desired number of cells in dishes and monitor cell growth by light microscope (see Note 12).

3. Use cells to perform any experiments as designed previously (i.e., transfect the cells with siRNA or vector containing a protein of interest; treat the cells with inhibitor(s) or any other chemicals). When cells are ready and have grown to desired confluence, transfer discs with cells to a new dish with warm media. Cells can be kept in incubator before freezing. This is helpful to avoid scratching and damaging cells on sapphire discs when transferring discs to specimen carrier for high-pressure freezing.

3.2. High-pressure freezing

• 1Prepare the Leica EM PACT2 with RTS according to the instruction manual. Operations are controlled through a touch screen on the front of EM PACT2. After switching on, the initialization screen followed by preparation screen will appear. Properly fill the hydraulic system with hydraulic fluid (e.g., methylcyclohexane), fill the Dewar via funnel with liquid nitrogen. The filling status of hydraulic fluid and liquid nitrogen are indicated on the preparation screen.
• 2In a 15 ml tube, prepare the cryoprotectant by mixing 0.2 g of ultra-low gelling temperature agarose, 0.182 g of mannitol and 200 μl of FBS in 10 ml DPBS. Agarose can be dissolved in DPBS at 60°C the day before using. Store at room temperature (RT). The next day, melt the agarose at 37°C and then add mannitol and FBS. Make up to 10 ml with DPBS. Keep the cryoprotectant in the tube at 37°C before and after each sample loading. For convenience, the cryoprotectant can be aliquoted into several 35-mm tissue culture dishes and kept at 37°C incubator.
• 3Always have water bath, incubator and stereo microscope with a warmed stage close at hand. Pre-warm loading tools including live cell carriers (1.5 mm inner diameter), rapid loader, forceps, RTS bayonet loading devices and bayonet pods (Figure 2B). Close the safety cover of RTS. Open valve by touching the valve symbol on Preparation screen to fill the liquid nitrogen bath.
• 4Touch “PRIME” button on the screen to move the pneumatic piston of the RTS to its right end position. Connect empty bayonet pod to a RTS loading device and turn clockwise to lock it. Open safety cover of RTS, place loading device into RTS and gently push loading device into position. Touch “Preload” button, the diamond locking screw is automatically turned clockwise and then turned back counterclockwise to open just enough to allow room for a flat specimen carrier. Both LED on the front of the RTS loading platform and the LED symbol on the touch screen turn green. The RTS is now ready for freezing a specimen.

Figure 2. High-pressure freezing-loading of samples and freezing.

(A) Loading sapphire disc containing monolayer cells into specimen carrier (filled with cryoprotectant) on the rapid loader. (B) During HPF, the specimen carrier and sapphire disc are inserted into the bayonet pod in liquid nitrogen. The arrow indicates the position of specimen carrier and sapphire disc in the pod. Note that sample carrier and sapphire disc remain together during subsequent steps through embedding in plastic.

The following specimen preparation procedures (steps 5 and 6) are performed on a warmed stage under stereo microscope.

• 5There are two alternative ways to prepare the rapid loader. First, place a specimen carrier upside down on the warmed stage. Gently pick up the specimen carrier into the rapid loader. Turn the rapid loader with carrier up again. Second, place a specimen carrier with empty recess, using fine forceps to pick up the specimen carrier and put it on recess of rapid loader. Open the recess the rapid loader gently with the fine forceps until carrier drops into the recess and is located correctly.
• 6Fill the specimen carrier with 5–10 μl cryoprotectant. Pick up sapphire disc with a cell monolayer and rinse the disc in the warmed cryoprotectant, and then insert the disc into specimen carrier (Figure 2A) (see Note 13). The sapphire disc/specimen carrier assembly is the sample transfer unit all the way through embedding into plastic. The carrier permits easy transfer without sample damage, step to step through much of the remaining protocol.
• 7Place rapid loader with specimen carrier and sapphire disc onto loading platform. The cell monolayer on the disc is facing to the right (see Note 14). Gently push the rapid loader in until you reach the end stop. Release the rapid loader and the high-pressure freezing procedure will be completed automatically in approximately 2.5 seconds. The rapid loader is released, and the specimen is ejected into the liquid nitrogen bath (Figure 2B). The process screen appears automatically to show the freezing parameters.
• 8To unload the specimen, all tools including wrenches and forceps must be precooled in liquid nitrogen before touching the frozen carrier. In the liquid nitrogen bath of EM PACT2, transfer pod into unloading station and disconnect bayonet loading device by gently pushing and turning counterclockwise. Use a wrench to turn counterclockwise to open the diamond screw in the bayonet pod. Pick up the specimen carrier using forceps and put it in container in liquid nitrogen bath. The specimens are now ready for the following freezing-substitution procedures.
• 9Touch “PREP” button and the screen changes back to preparation screen. The instrument is now ready for the next prime and preloading procedure.
• 10Repeat steps 4 to 9 as needed.
• 11When all samples are done, empty the dewar by draining liquid nitrogen into a storage dewar. As with all liquid nitrogen handling steps, insulated protective glove and protective eye wear should be worn (see Note 1). Activate bakeout cycle to dry all liquid nitrogen-cooled parts in the dewar. The bakeout cycle stops automatically.

3.3. Freeze-substitution

• 1Specimens are freeze-substituted using a Leica AFS unit (Leica Microsystems). Fill the dewar on AFS with liquid nitrogen. The filling level is indicated on main screen. As soon as dewar is full, stop. Precool chamber to −90°C.
• 2To prepare acetone containing 0.1% uranyl acetate / 0.5% glutaraldehyde, mix 50 μl of 10% uranyl acetate and 250 μl of 10% glutaraldehyde in 4700 μl of anhydrous acetone. Aliquot 500 μl of solution into pre-labeled 2 ml cryovials, and freeze under liquid nitrogen (see Note 15).
• 3To prepare acetone containing 0.1% uranyl acetate / 1% osmium tetroxide (OsO₄), mix 50 μl of 10% uranyl acetate and 400 μl of 12.5% OsO₄ in 4550 μl of anhydrous acetone (see Notes 2, 3 and 4). Aliquot the solution as described in Step 2. Keep these cryovials in Leica AFS chamber (−90°C).
• 4Use precooled forceps to transfer cold specimen carriers (Figure 2B) under liquid nitrogen into cryovials with frozen acetone containing 0.1% uranyl acetate / 0.5% glutaraldehyde. Close the vials with caps and keep them in liquid nitrogen.
• 5Transfer frozen specimen vials into −90°C chamber on Leica AFS unit. Place lid on top of chamber and turn lid clockwise to close lock. The properly mounted lid is indicated on the screen with the lid symbol above the dewar symbol. Specimens are freeze substituted at −90°C for 1–2 days.
• 6
  Transfer specimens into cryovials with acetone containing 0.1% uranyl acetate and 1% OsO₄, and perform the following program. The program can be created using AFS2 Program Editor Software.

  1. The specimens are warmed 3°C/h to −60°C and freeze substituted at −60°C for 8 h.
  2. The specimens are warmed 3°C/h to −30°C and freeze substituted at −30°C for 8 h.
  3. The specimens are warmed 3°C/h to 0°C.

As the vials warm from −90°C to 0°C, the freeze-sub cocktail will thaw, and the specimens will sink to the bottom.

• 7After the specimens are warmed to 0°C, remove the freeze substitution solution and wash the specimens 3 × 10 min with acetone. When removing solutions, always leave a little so the specimens do not dry out. Specimens are then warmed to RT.
• 8Use forceps to transfer specimens into 15 ml glass bottles filled with acetone. Make sure the cells on sapphire discs are facing up.

3.4. Infiltration and Embedding

1. Prepare 25%, 50%, 75% and 100% embedding medium by diluting it in anhydrous acetone.

2. Replace acetone with 25% embedding medium and irradiate the specimen bottles in microwave processor for 3 min at 250 Watts.

3. Remove the 25% embedding medium and replace with 50% embedding medium. Irradiate the specimen bottles in microwave processor for 3 min at 250 Watts.

4. Remove the 50% embedding medium and replace with 75% embedding medium. Irradiate the specimen bottles in microwave processor for 3 min at 250 Watts.

5. Remove the 75% embedding medium and cover the specimens with 100% embedding medium. Irradiate the specimen bottles in microwave processor for 3 min at 250 Watts.

6. Remove 100% embedding medium and replace with fresh 100% embedding medium. Irradiate the specimen bottles in microwave processor for 3 min at 250 Watts. Repeat this step twice.

7. When commercial microwave processor is not available, the infiltration procedures can be done at RT. Incubate the specimens in the glass bottles with 25%, 50%, 75% and 100% of embedding medium for at least 2 h, respectively. Then remove 100% embedding medium and replace with fresh 100% embedding medium and keep the specimens in fresh embedding medium overnight. Slow shaking is necessary to allow specimens to be well infiltrated. The following day, prepare new resin with DMP-30. Replace the old mixture with fresh resin. Specimens are now ready for the embedding procedure.

8. Transfer specimens from glass bottles into flat embedding mold with cells side up. Fill the mold with embedding medium (see Note 16). Cure the medium in an oven at 60°C for 24–48 h until it is hard. The blocks can be removed from the mold after it returns to RT (see Note 17).

9. Secure the block in specimen chunk in the microtome. Use a single-edge razor blade to remove excess resin surrounding specimen carrier. The carrier and sapphire disc are removed by repeated short freeze-thawing cycles in liquid nitrogen. After removal of disc and carrier, cells will stay in resin. The prior carbon coating of the sapphire disc before cell culture greatly facilitates the separation of cells from disc. (see Note 18). The blocks can be stored stable for years in dust-free conditions.

3.5. Sectioning and Imaging

1. Trimming the block for thin (50 nm) and semi-thick (300 nm) sections: Place the block back in specimen chunk in the microtome. Check monolayer cells under stereo microscope mounting on the ultramicrotome. Locate the area with cells of interest and mark it using razor blade. Trim the block so that the sectioning surface is a neat trapezoid shape, generally < 1 mm on a side. The most important trimming cuts are the bottom and the top of the block must be parallel.

2. Thin section (50 nm) transmission electron microscopy: Once trimmed, the block is mounted on an ultramicrotome. Sections are cut by a diamond knife and float off the edge onto water in the knife. After a ribbon of serial sections is formed, stop cutting. Flatten the sections with chloroform. Collect them on a grid and let it dry on a piece of filter paper. Sections are post-stained with uranyl acetate and lead citrate (see Step 3.6), and imaged with a FEI Tecnai TF20 intermediate-voltage electron microscope operated at 80 keV.

3. Semi-thick section (300 nm) electron tomography: the block is trimmed as in step 1. 300 nm semi-thick sections are cut using ultramicrotome and then collected onto slot grids. Sections are post-stained with uranyl acetate and lead citrate (see Step 3.6), and imaged with a FEI Tecnai TF30 intermediate-voltage electron microscope operated at 300 keV.

4. SBF-SEM: the full cell volumes can be obtained using a Zeiss Sigma VP scanning electron microscope equipped with a Gatan 3View SBF microtome system. The diamond equipped on the microtome cut the block in between imaging. Images are acquired using a backscattered electron detector. The image stacks are aligned to generate cell 3D data [16–19].

3.6. Post Sectioning Staining

1. Filter uranyl acetate and lead citrate solutions before use.

2. Prepare two petri dishes. Place an appropriate size piece of parafilm in each dish. In one dish, place several NaOH pellets around the edge of parafilm to make a CO₂ free environment.

3. Place one drop of uranyl acetate solution for each grid on first parafilm surface (without NaOH pellets). Float grid section side down on the drop and stain for 10 min in dark.

4. Carefully remove grid and rinse vigorously for 15 sec each in 2 changes of water.

5. Place one drop of lead citrate solution for each grid on second parafilm surface (with NaOH pellets). Float grid section side down on the drop and stain for 5 min.

6. Carefully remove grid and rinse vigorously for 15 sec each in 2 changes of water.

7. Blot dry and store the grids in a grid storage box.

3.7. Identify Golgi apparatus under electron microscope

In most mammalian cells, the highly organized Golgi apparatus is made up of Golgi stacks that consist of a series of flattened, membrane-bound cisternae. It is located in the cytoplasm near cell nucleus. In order to find the Golgi apparatus under an electron microscope, we recommend to first look at relatively low magnification in the general area of cell nucleus. Once the Golgi apparatus in a cell is located, switch to a higher magnification as needed and re-focus to get images.

4. Notes

1. Liquid nitrogen has extremely low temperature. Handling liquid nitrogen carelessly may cause cold burns. Insulated gloves and protective eyeware should be used.

2. OsO₄, glutaradehyde and uranyl acetate are very toxic. The resin components are toxic until completely polymerized. Always wear gloves and work in a fume hood.

3. Both OsO₄ and uranyl acetate are light sensitive. Keep these solutions in the dark and treat as potentially toxic. Osmium is volatile and should be used in a fume hood.

4. Uranyl acetate can degrade over time. The uranyl acetate solution should not be used if a cloudy precipitated forms.

5. Mix gently to avoid bubbles.

6. Aggressive mixing introduces air bubbles to the mixture. This will result in soft resin blocks after embedding.

7. Lead citrate strongly reacts with CO₂ and forms insoluble carbonate precipitates on stained sections. Using boiled water for making and rinsing solutions can eliminate CO2 and avoid the formation of precipitates.

8. The solution is cloudy before adding NaOH. Solution is clear after mixing well with NaOH.

9. The lead citrate solution should not be used if a cloudy precipitated forms.

10. Based on our experience, putting a small droplet of cell media on top of sapphire discs before adding all the media is helpful for preventing discs from floating.

11. Always handle the discs using very fine forceps and do not touch the coated surface.

12. We recommend preparing at least 2 sapphire discs for each experimental condition in case of cells loss or poor cell structure in some samples.

13. Be careful not to introduce any air bubbles while setting up the sample. During freezing, any air bubbles can cause the sapphire discs to shatter.

14. Handle the rapid loader gently to avoid falling of sapphire disc and specimen carrier.

15. Loose the caps of vials before freezing to avoid “pop” from trapped gas.

16. We recommend filling the mold slowly to avoid any air bubbles inside the embedding medium.

17. We usually use a single-edge razor blade to cut off the plastic mold.

18. Always allow the sapphire disc to be removed from the resin gently and leave the resin surface smooth and flat.