Here we describe a procedure for studying freeze-fractured plant tissues. High-pressure frozen leaf samples are freeze-fractured and double-layer coated, yielding well preserved frozen-hydrated samples that are imaged using the cryo-scanning electron microscope at high magnifications with minimal beam damage.
Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we describe a technique for studying the supramolecular organization of photosynthetic (thylakoid) membranes within leaf samples. This is achieved by high-pressure freezing of leaf tissues, freeze-fracturing, double-layer coating and finally cryo-SEM imaging. Use of the double-layer coating method allows acquiring high magnification (>100,000X) images with minimal beam damage to the frozen-hydrated samples as well as minimal charging effects. Using the described procedures we investigated the alterations in supramolecular distribution of photosystem and light-harvesting antenna protein complexes that take place during dehydration of the resurrection plant Craterostigma pumilum, in situ.
Oxygenic photosynthesis, originating in ancient cyanobacteria, was inherited by algae and land plants by endosymbiotic events that led to development of the chloroplast organelle. In all modern-day oxygenic phototrophs, photosynthetic electron transport and the generation of proton-motive force and reducing power are carried out within flattened sac-like vesicles termed 'thylakoid' membranes. These membranes house the protein complexes that carry out the light-driven reactions of photosynthesis and provide a medium for energy transduction. The thylakoid membranes of plants and (some) algae are differentiated into two distinct morphological domains: tightly appressed membrane regions called 'grana' and unstacked membranes that interconnect the grana, called 'stroma lamellae'1. Various freeze-fracture studies of plant and algal thylakoid membranes have been conducted, starting in the early 1970s. When freeze-fractured, membranes split along their hydrophobic core2, generating an exoplasmic face (EF) and a protoplasmic face (PF), depending on the cellular compartment which the half-membrane borders, as originally coined by Branton et al. in 19753. Plant and algal thylakoids have four different fracture faces: EFs, EFu, PFs and PFu, with 's' and 'u' denoting 'stacked' and 'unstacked' membrane regions, respectively. The membrane protein complexes, which are not split or broken, have the tendency to remain with either the E or P side of the membrane. The initial observations that the different fracture faces of the thylakoids contain particles of different sizes and densities4, and the numerous investigations that followed, led to identification and correlation between the observed particles and the membrane protein complexes that carry out the light reactions5-13 (see also reviews14,15).
Freeze-fracture experiments of thylakoid membranes are typically carried out on preparations of chloroplasts or isolated thylakoid membranes (but see16,17), at the risk of any alteration in structural and/or supramolecular organization that may occur during the isolation procedure. Following fracture, replicas are prepared by evaporation of platinum/carbon (Pt/C), then by a thick layer of carbon (C), and finally digestion of the biological material18. Replicas are visualized by transmission electron microscopy (TEM). The traditional freeze-fracture-replica technique continues to serve as an important tool for studying the supramolecular organization of photosynthetic membranes and their adaption to different, e.g., light, conditions19-23.
In our recent study of the homoiochlorophyllous resurrection plant Craterostigma pumilum24, we aimed to investigate the changes in the supramolecular organization of thylakoid membranes, as well as in overall cellular organization, during dehydration and rehydration. The uniqueness of homoiochlorophyllous resurrection species is that they are able to survive conditions of desiccation in their vegetative tissues (leaves), while retaining their photosynthetic apparatus. Once water is available, these plants recover and resume photosynthetic activity within hours to a few days25. For this study, cryo-scanning EM (SEM) imaging of freeze-fractured leaf samples was combined with high-pressure freezing for sample cryo-immobilization. These procedures provide a means to visualize frozen-hydrated biological samples at a state close to their native state26. One main benefit is that samples are examined directly after freeze-fracture and coating with no successive steps. This is particularly relevant to the investigation of plants at different relative water contents (RWC), as their hydration state is maintained during preparation. However, one critical disadvantage is that frozen-hydrated samples may suffer from beam damage during imaging, especially when scanned at high magnifications, required for accurate measurement of the size of photosynthetic complexes. To overcome this, a method called 'double-layer coating' (DLC)27,28 combined with specific cryo-SEM imaging conditions were utilized. These resulted in samples that are significantly less beam-sensitive and allowed for the elucidation of valuable information on photosynthetic protein supramolecular organization and other cellular constituents of the resurrection plant C. pumilum at high magnifications in situ.
1. Cryo-fixation of Leaf Tissues by High-pressure Freezing
Note: This section describes how to carry out high-pressure freezing of leaf tissues for a freeze-fracture experiment. For considerations related to plant samples see29. This can be adapted for other types of tissues or samples with some modification.
2. Freeze-fracture and Double-layer Coating27,28
3. Cryo-scanning Electron Microscopy
4. Image Analysis
Note: This section describes a short procedure for segmentation of membrane particles from freeze-fracture SEM images using the Fiji31 open-source package. Similar results can be obtained with other image analysis software.
Figure 1 shows cryo-SEM images of platelets containing high-pressure frozen, freeze-fractured Craterostigma pumilum leaf pieces. In some samples, large regions of fractured cells are obtained (Figure 1A). In others, the leaf piece stays tightly bound to the upper disc and is knocked off along with it (Figure 1B). However, even in the second case, some leaf tissue may remain attached to the knife grooves on the platelet (Figure 1B, arrowheads) and a fair amount of data can still be collected by imaging the leaf "remnants", provided they were fractured. After successful fracture is found, low magnification images of cells are acquired to identify regions of interest, in this case chloroplasts (Figure 2).
Higher-magnification images of thylakoid membranes in C. pumilum leaves are shown in Figures 3 and 4. The four different fracture faces, EFs, EFu, PFs and PFu, can be distinguished (Figure 3). Photosystem II (PSII), which is the largest protein complex found in the membranes, is located in both the EFs (grana) and EFu (stroma lamellae). At hydrated conditions, PSII density is ~3 times higher in EFs than in EFu (~1,550 complexes µm-2 vs. ~550 complexes µm-2, Figure 3). This is one basis for differentiating between these two continuous faces. Another is the difference in their background. While the EFs background appears smooth, the EFu face is rough and contains holes that are the footprints of detached PSI complexes, which fracture to the complementary face, the PFu10 (Figure 3). An example for the segmentation of PSII complexes from the EFs face of the thylakoid membrane is shown in Figure 5.
During dehydration of C. pumilum, the density of PSII in the grana membranes (EFs) gradually decreases, reaching roughly half (~700 complexes µm-2 at ~15% RWC, Figure 4C) of their density at hydrated conditions (100% RWC, Figure 4A). Notably, upon further dehydration, to 5 – 10% RWC, PSII complexes organize into rows and arrays (arrows, Figures 4D and 4E). For complete dataset, see24. Formation of such PSII arrays necessitates that some of their bound antenna complexes, LHCII, detach from PSII. An example for LHCII complexes organized into rows in the PFs that would be complementary to organized PSII complexes (in the EFs) is shown in Figure 4E (arrowhead). PSII complexes in the arrays, as well as the detached LHCII, are likely to be in a photochemically quenched state, serving a photo-protective role. These structural rearrangements are part of the mechanisms utilized by the resurrection plant C. pumilum to protect itself in the dehydrated state24.
Figure 1. Low Magnification Cryo-SEM Images of Platelets with Freeze-fractured C. pumilum Leaf Pieces. (A) A large region of fractured cells (dashed outline) of a C. pumilum leaf. (B) The leaf piece was knocked off with the top platelet, but some leaf tissue remained attached to the knife marks at the bottom platelet (arrowheads mark some of these). The regions surrounding the leaf pieces are frozen 1-hexadecene (asterisks). Scale bars: 200 µm. Please click here to view a larger version of this figure.
Figure 2. Zooming into Chloroplasts. (A) A group of fractured photosynthetic (mesophyll) cells (asterisks) of hydrated leaf tissue. Most of the cell volume is comprised of a large central vacuole, with all cytoplasmic constituents surrounding the vacuole in a narrow strip. (B) Two neighboring fractured cells – the cell wall (cw) marks the border between the cells. Five chloroplasts (c) are apparent in the image, only one of which has been fractured (fractured plane marked by the arrowhead). (C and D) Examples for single fractured chloroplasts from hydrated (C) and dehydrated (D, ~15% relative water content [RWC]) plants. The small white dots (e.g., arrowheads) are the membrane protein complexes found within the thylakoid membranes of the chloroplast. Note the massive vesiculation surrounding the chloroplast found in the dehydrated leaf (D). Scale bars: 10 µm (A); 1 µm (B); 200 nm (C and D). Please click here to view a larger version of this figure.
Figure 3. The Supramolecular Organization of Thylakoid Membranes within Plant Tissues. A fractured chloroplast in which the different thylakoid fracture faces can be seen: the exoplasmic fracture faces (EF) of both stacked (EFs) and unstacked (EFu) membrane regions contain photosystem II (PSII) complexes; The protoplasmic fracture face of stacked regions (PFs) contains the peripheral light-harvesting antenna complexes of PSII, LHCII; Photosystem I (PSI) and ATP synthase fracture to the protoplasmic fracture face of unstacked regions (PFu); Cytochrome b6f fractures to both PFs and PFu. Image was scanned at a magnification of 203 kX (horizontal field of view = 1.45 µm). Scale bar: 100 nm. Please click here to view a larger version of this figure.
Figure 4. Changes in Photosystem II Organization during Dehydration of C. pumilum. EFs faces of thylakoid membranes of plants at different RWC: (A) 100%, (B) ~40%, (C) ~15%, and (D and E) 5 – 10%. During dehydration, the density of PSII in the EFs face gradually decreases from ~1,500 complexes µm-2 (A) to ~700 complexes µm-2 (~15% RWC, C). Notably, at drier conditions (5 – 10% RWC), some PSII complexes organize into rows and arrays (D and E, arrows). The arrowhead (E) marks the PFs face, with LHCII antenna appearing to also be organized in rows, in between which PSII rows would be found in the complementary EFs. For additional information on this data see24. Scale bars: 100 nm. Please click here to view a larger version of this figure.
Figure 5. An Example for Segmentation of Photosystem II Particles. Using the Fiji open-source package, protein complexes can be segmented from the images with a few steps (complete detailed procedure found in part 4 of the Protocol section). Images (A, original image) are blurred with the Gaussian filter (B); the image is thresholded using an auto local threshold tool (C); the image is inverted and the watershed algorithm is applied in order to separate objects found in close contact (D); a region of interest is selected and the outside is cleared (E); particles (within a user-defined size range) are selected using the particle analyzer function. The resulting mask is shown in (F). Particles are added to the ROI Manager and overlaid onto the original image (G) to check for errors or for faulty or missed particles. The list of ROIs can be manually edited, replacing, deleting or adding new ROIs, as appropriate. The final user edited selection is shown in (H). Scale bar: 200 nm. Please click here to view a larger version of this figure.
The technique described in this paper allows investigation of freeze-fractured membranes within the context of well-preserved high-pressure frozen plant tissues by cryo-scanning electron microscopy. The major advantage of using these procedures is that sample preparation is purely physical; no steps involving chemicals or dehydration are necessary. Thus, it allows studying biological structures at a near-native state26,32. The benefit of using leaf tissues is that one can obtain information on the overall cellular organization, as well as study specific membranes, such as the thylakoid membranes, within their native physiological context, i.e., within the chloroplast. Another advantage, imperative to studying C. pumilum plants at different relative water contents (RWC), is that their hydration state is not altered during preparation. This is as opposed to utilizing isolated chloroplasts or thylakoid membranes as the starting material for a freeze-fracture experiment. In the latter, one also has to take into account that some structural and/or supramolecular alterations likely take place during organelle/membrane isolation.
Working with plant tissue samples, which are relatively thick [>50 – 100 µm, and can be significantly thicker depending on the type of tissue and species], dictates the use of high-pressure freezing (HPF) for sample vitrification. Application of high pressure [2,100 bar], at which the melting temperature of water is about −22 °C, affects the hydrogen bond network of the water such that it slows down the rate of ice crystal formation. Therefore, the chance of obtaining a vitrified sample is high even at relatively slow cooling rates. HPF is currently the only method available for vitrification of thick, yet not exceeding 200 µm, samples. A critical step for HPF of plant tissues is infiltration of their intercellular air spaces with an inert "space filler" prior to freezing, to prevent tissue collapse under the high pressure (see reviews29,33). In the case of C. pumilum leaves, whose thickness can be up to 1 mm, the leaves also have to be trimmed or thinned prior to HPF.
Essentially, the technique of replica preparation18 can also be applied to freeze-fractured leaf tissues. However, in our experience, sufficient yield of intact fractured areas was not obtained, since replicas often fragment to small pieces. Moreover, searching for a specific region of interest such as chloroplasts can be impractical within fragmented replicas of plant tissues. This is mainly because mesophyll (photosynthetic) cells are comprised of a large central vacuole surrounded by a narrow cytoplasmic strip (Figure 2A), which includes the nucleus, chloroplasts and all other organelles. Thus, chloroplasts represent only a very small fraction of the total fractured area of mesophyll tissues. Cryo-SEM imaging of freeze-fractured samples offers a solution to this problem since samples are directly visualized following fracture (and coating/shadowing). However, this method suffers from the limitation that frozen-hydrated samples are beam-sensitive and thus scanning at high magnifications readily damages the sample on the first scan. Double-layer coating (DLC)27,28 offers a solution to this serious drawback.
In DLC, fractured samples are coated in a way similar to how they are for replica preparation. First, a thin layer (1 – 3 nm) of Pt/C is evaporated onto the sample, providing contrast and conductivity. This is followed by a thicker (5 – 10 nm), protective layer of carbon. DLC combined with a high accelerating voltage (10 kV) was used to image samples using the backscattered electron (BSE) signal28. The BSE signal, originating from the platinum layer allows obtaining surface information through the carbon and, possibly, water vapor contaminant layer, which are practically transparent to the BSE at high acceleration voltages (water contamination accumulates in experiments in which a vacuum-cryo transfer unit is not used and the fractured surface is exposed to the atmosphere, even if this is for a fraction of a second). In addition, imaging using the BSE signal minimized the problem of charging effects which are apparent in secondary electron (SE) detection27. With the microscope setup described here, imaging of the SE signal with the In-lens SE detector at 10 kV yielded information equivalent to that obtained when samples were coated solely with 2 – 3 nm of Pt/C and imaged at low accelerating voltages. The difference is that samples prepared by DLC and imaged as described were considerably less beam-sensitive. This enabled us to scan them at high magnifications and in many cases even repeatedly, and allowed obtaining valuable information on photosynthetic protein supramolecular organization and other cellular constituents (Figures 2 – 4)24.
Finally, the procedures described here can be modified to study the structure and/or membrane protein organization in other types of samples. We have successfully employed the technique to study the supramolecular organization of photosynthetic membranes in cyanobacterial and algal cells (Shperberg-Avni et al. unpublished data). The main consideration is finding the balance between having as thin a sample as possible, in order to increase the chance for vitrification, while keeping sample intactness. For different types of samples, which can be suspensions or animal/plant tissues, different packaging should be used (e.g., in terms of the type of platelets utilized for high-pressure freezing). Once successful cryo-immobilization is achieved, the combination of freeze-fracture, DLC and cryo-SEM imaging provides an excellent means for obtaining information on membrane protein organization at high magnification with minimal beam damage.
The authors have nothing to disclose.
We thank Andres Kaech (University of Zurich) for his helpful advice on scanning electron microscopy imaging. This work was supported by the United States-Israel Binational Agricultural Research and Development Fund (grant no. US-4334-10, Z.R.), the Israel Science Foundation (grant no. 1034/12, Z.R.), and the Human Frontier Science Program (RGP0005/2013, Z.R.). The electron microscopy studies were conducted at the Irving and Cherna Moskowitz Center for Nano and Bio-Nano Imaging at the Weizmann Institute of Science.
ethanol abs | Bio-Lab | 052505 | |
isopropanol | Bio-Lab | 162605 | |
1-hexadecene | Sigma-Aldrich | H7009 | |
0.1/0.2 platelets | Engineering Office M. Wohlwend GmbH, Switzerland | 241 | Platelets are of 3-mm diameter and 0.5-mm-thick (Type A) with 0.1/0.2-mm-deep cavities (of diamater 2 mm). Similar platelets can be obtained from Leica Microsystems. |
high-precision-grade tweezers | Electron Microscopy Sciences | 72706-01 | Dumont (Switzerland) Durostar style #5 tweezers; Can be substituted with other high-precision tweezers. |
high-pressure freezing machine | Bal-Tec | HPM 010 | High-pressure freezing alternatives: 1. HPF Compact 02, Wohlwend GmbH; 2. HPM 010, RMC Boeckeler; 3. EM PACT2, Leica Microsystems; 4. EM HPM 100, Leica Microsystems; 5. EM ICE, Leica Microsystems. |
freeze-fracture system | Leica Microsystems | EM BAF 060 | |
cryo preparation loading stage | Leica Microsystems | 16770228 | |
specimen holder for univeral freeze fracturing | Leica Microsystems | 16LZ04746VN | Clamp holder for specimen carriers of diameter 3 mm |
vacuum cryo-transfer shuttle | Leica Microsystems | EM VCT 100 | |
scanning electron microscope | Zeiss | Ultra 055 | |
cryo SEM stage | Leica Microsystems | 16770299905 | |
image acquisiton software | SmartSEM, Carl Zeiss Microscopy GmbH | ||
image analysis software | Fiji/Image J, National Institute of Health | http://fiji.sc/Fiji |