Abstract
Chloroplasts, as well as other, non-photosynthetic types of plastid, are characteristic structures within plant cells. They are relatively large organelles (typically 1–5 μm in diameter), and so can readily be analysed by electron microscopy. Chloroplast structure is remarkably complex, comprising at least six distinct sub-organellar compartments, and is sensitive to developmental changes, environmental effects, and genetic lesions. Transmission electron microscopy (TEM), therefore, represents a powerful technique for monitoring the effects of various changing parameters or treatments on the development and differentiation of these important organelles. We describe a method for the analysis of Arabidopsis plant material by TEM, primarily for the assessment of plastid ultrastructure.
Key words
1 Introduction
Plastids are a functionally and structurally diverse group of related organelles, including photosynthetic chloroplasts and a variety of non-photosynthetic variants such as leucoplasts, amyloplasts, and chromoplasts (1, 2). They are characteristic components of plant cells and have been subjected to extensive ultrastructural analysis by electron microscopy (EM) over many years. Such analysis is invaluable, as organellar structure is influenced by a variety of factors including developmental, environmental, and genetic parameters (3–5). Thus, ultrastructural analysis of plastids enables the consequences of various treatments and conditions for organelle structure and development to be analysed and quantified.
Electron microscopy has many variants; even a focused approach to a single specimen has many options that may be explored. We have aimed to describe procedures based on access to the expertise and not insignificant physical resources of an established electron microscopy laboratory. We provide a basic methodology for the collection, preservation (fixation), and embedding of the sample. Brief descriptions of the processes of ultramicrotomy and section staining, and use of the transmission electron microscope are also provided, but detailed instructions for these are beyond the scope of this text and are described in many other publications, for example, Kuo (6). The described techniques are those we have successfully employed, and they should be readily reproducible in any electron microscopy laboratory. By restricting ourselves to these conventional methods, we avoid the need for specialised preparative apparatus, such as that needed for high pressure freezing and freeze substitution techniques as utilised by Pfeiffer and Krupinska (7). It should be noted that the described methods are far from exhaustive and indeed are offered as a starting point from which modifications may be made to suit more “local” applications.
2 Materials
2.1 Major Equipment
All of the following items are essential to these procedures. However, similar items should already be present in any biological sciences EM laboratory. This summary is based upon what we use and does not imply superiority of one brand or manufacturer over another.
-
1.
A ducted fume cupboard, suitable for handling flammable and carcinogenic materials (i.e., it should be spark proof and not of the portable, re-circulating variety).
-
2.
Spark-proof refrigerator or cold room, set at 4°C.
-
3.
Stereo dissecting microscope (e.g., Olympus SZ51, Olympus UK Ltd., Southend-on-Sea, Essex, UK).
-
4.
Ultramicrotome (e.g., Leica EM UC7, Leica Microsystems (UK) Ltd., Milton Keynes, UK).
-
5.
Light Microscope (e.g., Olympus BX Series, Olympus UK Ltd.).
-
6.
Transmission electron microscope. We have a JEOL 1400 (JEOL (UK) Ltd., Welwyn Garden City, UK).
2.2 Minor Equipment
-
1.
Fine-point stainless steel forceps (e.g., Dumont tweezers 4, T5288, Agar Scientific Ltd., Stansted, UK).
-
2.
Reverse forceps (e.g., Dumont tweezers, NOC, T5071, Agar Scientific Ltd.).
-
3.
Fine artists’ paint brush (Size 00) (e.g., G3444, Agar Scientific Ltd.).
-
4.
Variable-speed rotary mixer (e.g., R060, TAAB Laboratories Equipment Ltd., Aldermaston, UK) or a variable-speed roller mixer (e.g., 444–1067, VWR International Ltd., Lutterworth, UK).
-
5.
Portable balance; range 0–200 g, with accuracy to two decimal places (e.g., Fisherbrand SG202, Fisher Scientific, Loughborough, UK).
-
6.
Variable-speed magnetic stirrer with stirring bars (e.g., Hanna HI-200M, Fisher Scientific).
-
7.
Embedding oven, chest type (e.g., EO62 or E100, TAAB Laboratories Equipment Ltd.).
-
8.
Embedding capsule press (e.g., C213, TAAB Laboratories Equipment Ltd.); this item is not essential but it is extremely useful.
-
9.
Small bench vice (e.g., “Vacu-Vice,” T576, Agar Scientific Ltd.).
-
10.
Small (6 in.) “Junior” hacksaw.
-
11.
Hotplate (e.g., Stuart SD160, VWR International Ltd.).
2.3 Consumables and Small Items
-
1.
Personal protective equipment, including disposable protective gloves and laboratory coat.
-
2.
Locally approved incineration bins for disposal of solid waste, resin-contaminated material, and sharps.
-
3.
Specimen tubes, 2 mL (T308-2, Simport, Beloeil, Canada).
-
4.
Disposable graduated plastic transfer pipettes, 7 mL (612–1681, VWR International Ltd.).
-
5.
Ordinary glass microscope slides (plain), or white tile.
-
6.
Razor blades, single or double edged (e.g., T585, Agar Scientific Ltd.).
-
7.
Pipettors and tips, to handle volumes ranging from 5 μL to 1 mL (e.g., Finnpipette or Gilson Pipetman).
-
8.
Polyethylene graduated containers with caps (e.g., G332 and G333, Agar Scientific Ltd.).
-
9.
Polyethylene flat-ended embedding capsules (for modified Spurr’s resin or other epoxy resins) (e.g., G3744, Agar Scientific Ltd.).
-
10.
Petri dish, 9-cm diameter, glass (or Conway Dish if available).
-
11.
Plastic sealing film (e.g., Parafilm).
-
12.
Syringe filters, 0.2 μm (e.g., Acrodisc, Pall Corporation, MI, USA).
-
13.
Electron microscope grid storage box (e.g., G276, Agar Scientific Ltd.).
2.4 Chemistry (see Note 1)
The following is a list of chemicals that are required for the described techniques. We assume that the reader has access to a distilled water supply.
2.4.1 Tissue Fixation
-
1.
Sörensen’s phosphate buffer (SP buffer), comprising disodium hydrogen orthophosphate and sodium dihydrogen orthophosphate (see Subheading 2.5.1).
-
2.
Glutaraldehyde, 25% (w/v) aqueous solution, EM grade. Toxic and harmful to the environment.
-
3.
Paraformaldehyde (prilled), EM grade. Harmful.
-
4.
Osmium tetroxide, 2% (w/v) aqueous solution. Toxic and corrosive.
-
5.
Milk powder.
-
6.
Potassium ferricyanide. Irritant.
-
7.
Uranyl acetate (also used in section staining). Radioactive, very toxic, and may have cumulative effects.
-
8.
Sodium hydroxide, 1 N (1 M), carbonate-free aqueous solution. Corrosive.
2.4.2 Tissue Dehydration and Embedding
-
1.
Ethanol, analytical grade. This is used both pure and diluted (in distilled water) at the following concentrations: 30, 50, 70, and 90% (v/v). Highly flammable.
-
2.
1,2-Epoxypropane (pseudonym, propylene oxide; PO). Toxic, suspected carcinogen, and highly flammable.
-
3.
Epoxy resin embedding medium. For example, modified Spurr’s resin, comprising: ERL 4221 (irritant); diglycidylether of polypropylene glycol (DER 736, irritant); nonenyl succinic anhydride (NSA, irritant); dimethylaminoethanol (DMAE, flammable and irritant).
2.4.3 Sample Remounting and Section Staining
-
1.
High-strength epoxy resin adhesive (e.g., Araldite, Huntsman Advanced Materials Ltd., Cambridge, UK). Irritant and dangerous to the environment.
-
2.
Reusable pressure adhesive (e.g., Blu-Tack, Bostik Ltd., Leicester, UK).
-
3.
Lead nitrate (to make Reynolds lead citrate). Toxic and dangerous to the environment.
-
4.
Trisodium citrate (to make Reynolds lead citrate).
2.5 Working Buffers and Reagents (see Note 1)
2.5.1 Sörensen’s Phosphate Buffer (0.2 M, pH 7.2) (8)
Sörensen’s phosphate buffer (SP buffer) is a mixture of mono- and dibasic sodium phosphates. Prepare a 0.2 M disodium hydrogen orthophosphate solution (Solution A) as follows: dissolve 35.61 g of Na2HPO4⋅2H2O (or 53.65 g of Na2HPO4·7H2O or 71.64 g of Na2HPO4⋅12H2O, depending on availability) in 1 L of distilled water. Next, prepare a 0.2 M sodium dihydrogen orthophosphate solution (Solution B) as follows: dissolve 27.6 g of NaH2PO4·H2O (or 31.21 g of NaH2PO4⋅2H2O) in 1 L of distilled water.
To make the 0.2 M SP buffer stock solution, mix 36 mL of Solution A with 14 mL of Solution B; the pH of the final solution should be 7.2, which should be verified. This stock solution is used to make the primary and secondary fixation solutions (see Subheading 2.5.3). A further dilution of the stock is also required: 0.1 M SP buffer is used for washing steps (see Subheading 3.2).
2.5.2 Depolymerised Paraformaldehyde Solution (10% Formaldehyde)
In a fume cupboard, dissolve 10.0 g of prilled paraformaldehyde in 90 mL of distilled water. Warm the mixture to approximately 60°C and stir constantly to ensure that the paraformaldehyde dissolves. If the solution is not completely clear, add, drop-wise, 1 N sodium hydroxide until clarity is achieved. The solution should then be made up to 100.0 mL with distilled water. Ideally, this should be prepared immediately before use (9), though it may be stored frozen for some months. If the solution exhibits any opalescence or traces of flocculate material, it should not be used.
2.5.3 Primary, Secondary, and Tertiary Fixatives
-
1.
Primary fixative. Add 4.0 mL of freshly depolymerised 10% (w/v) paraformaldehyde solution (see Subheading 2.5.2) to 5.0 mL of 0.2 M SP buffer. To this mixture, add 1.0 mL of 25% (w/v) glutaraldehyde solution. The final working solution is 4.0% (w/v) formaldehyde, 2.5% (w/v) glutaraldehyde, in 0.1 M SP buffer (10) (see Note 2).
-
2.
Secondary fixative. To 1.0 mL of 2% (w/v) aqueous osmium tetroxide (see Notes 3 and 4), add 1.0 mL of 0.2 M SP buffer, and to this add 30.0 mg of potassium ferricyanide. The final working solution is 1% (w/v) osmium tetroxide in 0.1 M SP buffer containing 1.5% (w/v) potassium ferricyanide.
-
3.
Tertiary fixative. To 1.0 mL of distilled water, add 20.0 mg of uranyl acetate (see Note 5). The final working solution is 2% (w/v) aqueous uranyl acetate. This solution may also be used as an electron-dense stain for ultrathin sections (see Subheading 3.4.1).
2.5.4 Modified Spurr’s Low-Viscosity Resin (11) (see Notes 6–8)
The following procedure makes approximately 30.0 mL of “standard” hardness resin. Using disposable 7-mL graduated plastic transfer pipettes (see Notes 9 and 10), weigh into a disposable plastic container (see Subheading 2.3, item 8) 12.3 g of ERL 4221, 4.3 g of DER 736 (see Note 11), and 17.7 g of NSA (see Subheading 2.4.2). Add a small magnetic stirring bar and thoroughly mix for 5 min on a magnetic stirrer. Finally, add 0.3 g of DMAE accelerator and mix for a further 5 min. Avoid incorporating air bubbles into the mixture. DMAE and NSA should never be mixed together alone as this may result in a strong exothermic reaction.
Test the quality of the resin as follows: fill a polyethylene flat-ended embedding capsule (see Subheading 2.3, item 9) with the resin mixture and polymerise for at least 8 h at 70°C. Remove the polymerised block from the capsule and check the hardness and integrity of the resulting block. This is best judged by using a single-edged razor blade to trim a piece of resin from the block; it should not be soft or brittle.
2.5.5 Reynolds Lead Citrate (12) (see Note 12)
Boil and allow to cool 100 mL of distilled water (see Note 13). Weigh 1.33 g lead nitrate (Pb(NO3)2) and 1.76 g trisodium citrate (Na3(C6H5O7)⋅2H2O) into a 50-mL volumetric flask. Add 30 mL of the boiled distilled water to the flask. Close the flask with a stopper and then shake vigorously for at least 1 min. Leave the mixture to stand, with occasional shaking, to complete the conversion to lead citrate. The mixture will become “milky” in appearance and should contain no solid material.
After 30 min, add 8.0 mL of 1 N sodium hydroxide solution (13); the mixture should “clear” quickly. Dilute to 50 mL with additional freshly boiled, distilled water, and mix by inversion. The resulting solution should have a pH of 12 (±0.1) and should be stored at 4°C in a tightly sealed bottle. Before use, allow to reach room temperature and filter through a 0.2-μm syringe filter to remove any particulate matter.
3 Methods
The entire procedure, from start to finish, will take approximately four days; therefore, sample collection should ideally commence at the beginning of the week.
To process two genotypes (i.e., at least six cotyledons or leaf sections) in 2.0 mL specimen tubes for transmission electron microscopy (TEM) requires approximately 21.0 mL of distilled water, 5.0 mL of 0.2 M SP buffer, 0.8 mL of 10% freshly depolymerised paraformaldehyde (formaldehyde), 0.2 mL of 25% glutaraldehyde, 1.0 mL of 2% osmium tetroxide, 30.0 mg of potassium ferricyanide, 12.0 mL of analytical grade ethanol (including that used for graded dehydration steps), and 8.0 mL of 1,2-epoxypropane (including that used for graded infiltration steps). In practice, larger “stock” volumes of each of the above should be available. Approximately 30.0 mL of freshly mixed modified Spurr’s resin should be made.
3.1 Collection of Samples
3.1.1 Standardisation
When working with young seedlings, one may choose to analyse cotyledons, true leaves, or non-photosynthetic tissues such as the roots. With very young/small specimens, it may prove optimal to excise and process whole cotyledons or even process the entire seedling. It should be normal practice to take samples from seedlings or plants of equivalent age and which have been grown under identical conditions. When analysing true leaves, it is important to select leaves of an equivalent developmental stage.
3.1.2 Practice
When using plate-grown samples, identify and use seedlings whose cotyledons/leaves are not in contact with the growth medium. Cotyledons and small leaves may be excised from the seedling at their base with a razor blade. When possible, it is our practice to also cut away the top 30–50% of the cotyledon or leaf at 90° to the central vein (see Fig. 1) (see Note 14). The retained portion is immediately immersed in the buffered primary fixative solution (see Subheading 3.2.1). When working with true leaves, a slice (maximum 3 × 5 mm) is dissected from the central portion of the leaf lamina, again at 90° to the central vein. When working with roots, ∼2–3-mm-long sections approximately 3 mm back from the root apex should be excised for analysis. When working with dark-grown seedlings in order to study etioplast ultrastructure, tissue collection and initial fixation should be conducted under green safe-light conditions. Cotyledons from etiolated plants are collected by harvesting intact apical regions approximately 2–3 mm in length. Particularly small, entire seedlings may be fixed without dissection in primary fixative. These may then be washed and dissected before the secondary fixation stage, under normal lighting.
It is normal to retain only one cotyledon, leaf or root specimen per plant, in order that one can be sure that each corresponds to a separate individual. Several (at least six) unique specimens should be collected for processing, per sample (i.e., per genotype or condition), although usually only three will be sectioned, observed, and recorded.
3.2 Tissue Processing for TEM (see Note 1)
With the exception of the refrigerated steps, all of the following should be conducted in a suitable ducted fume cupboard at room temperature. Materials stored in a refrigerator should be allowed to reach room temperature before opening to avoid the possible formation of condensation.
The following procedure assumes the use of 2-mL screw-top specimen tubes (see Subheading 2.3, item 3). Alternative containers (e.g., 7-mL screw-top, flat-bottomed, glass vials) may be substituted with appropriate reagent volume adjustments (see Note 15). A minimum of 1.0 mL of each reagent should be used in all steps. The samples should be regularly agitated; this may be done intermittently by hand, or (preferably) constantly on a rotary or roller mixer.
3.2.1 Day 1
-
1.
For each sample (i.e., for each plant genotype or condition to be analysed), dispense at least 1.0 mL of freshly prepared primary fixative (see Subheading 2.5.3) into a labelled specimen tube.
-
2.
Transfer the biological material to be sampled to a glass microscope slide (or tile). To avoid physical damage, do not manipulate the area of tissue to be examined. Use a single-edged razor blade to excise the required tissue specimen, as described in Subheading 3.1.2, and immerse it immediately in the fixative solution (see Note 16). Repeat as many times as required with additional plants.
-
3.
Allow the samples to fix for ∼3–4 h at room temperature following collection of the last sample (see Note 17). After this time, move the vials to a refrigerator (ensure that the samples are still in contact with the fixative) and leave overnight (see Note 18).
3.2.2 Day 2
-
4.
Remove the samples from the refrigerator and allow them to reach room temperature.
-
5.
Using a disposable plastic transfer pipette, remove the fixative solution and replace it with 0.1 M SP buffer (see Notes 19–21). Wash for 30 min.
-
6.
Repeat the last step two more times. The waste fixative and washes must be disposed of appropriately.
-
7.
Remove the final wash and replace it with 1.0 mL of secondary fixative (see Subheading 2.5.3) (see Note 3). Allow the samples to fix for 1–1.5 h at room temperature.
-
8.
Thoroughly wash the samples (now noticeably dark greenish-black) once for 15 min with 0.1 M SP buffer, and then three times (for 30 min each time) with distilled water (see Note 22). The waste fixative and washes must be disposed of appropriately.
-
9.
Remove the final wash and replace it with 1.0 mL of tertiary fixative (see Subheading 2.5.3) (see Note 5). The samples should now be kept in the dark for 1 h; agitate them regularly.
-
10.
Wash the samples three times (30 min each time) in distilled water. The waste uranyl acetate and washes must be disposed of appropriately.
-
11.
After removing the final wash, commence dehydration of the samples using a graded ethanol/distilled water series as follows (in order of increasing ethanol concentration): 30, 50, and 70% ethanol, for 30 min at each step.
-
12.
Replace the 70% ethanol with fresh 70% ethanol, transfer the samples to a refrigerator, and leave overnight.
3.2.3 Day 3
-
13.
Remove the samples from the refrigerator and allow them to reach room temperature.
-
14.
Conduct further dehydration steps using (in order) 90% ethanol and 100% analytical grade ethanol for 30 min each (see Note 23). The analytical grade ethanol step should be repeated twice more.
-
15.
During step 14, prepare the required volume (dependent on the number of samples) of modified Spurr’s resin (see Subheading 2.5.4) to be used in the following infiltration and embedding steps.
-
16.
Exchange the ethanol for 1, 2-epoxypropane (PO) and incubate for 10 min, and then repeat. PO is a toxic, potentially carcinogenic, and highly flammable solvent (see Note 24). Do not allow the samples to be exposed to air; leave a minimal residual amount of the previous solution during exchanges.
-
17.
Infiltrate the samples with the modified Spurr’s resin by immersion in progressively increasing concentrations of resin according to the following series (see Note 25):
-
(a)
3 parts PO to 1 part resin, 60–90 min;
-
(b)
1 part PO to 1 part resin, 60–90 min;
-
(c)
1 part PO to 3 parts resin, 60–90 min;
-
(d)
100% resin, 30 min.
-
(a)
-
18.
Replace the resin with fresh 100% resin and leave on the mixer overnight.
3.2.4 Day 4
-
19.
Replace the overnight resin with fresh 100% resin and leave on the mixer at room temperature for a further 3 h.
-
20.
Obtain the required number of polyethylene embedding capsules (see Subheading 2.3, item 9). Each capsule should include a label indicating the identity of the sample (see Note 26).
-
21.
Using a modified 7-mL disposable plastic transfer pipette, collect an individual leaf specimen from the processing container and transfer it (with a small volume of resin) into a separate capsule, i.e., one piece per capsule (see Note 27). Repeat this for each specimen.
-
22.
Completely fill each capsule with fresh 100% resin and close the lid (see Notes 7 and 28).
-
23.
Leave the loaded capsules for an hour or two and then transfer them into a pre-heated (70°C) embedding oven for at least 8 h to polymerise the resin.
-
24.
Proceed to Subheading 3.3.
3.3 Remounting (see Fig. 2)
For sections to be taken at a specific orientation from the tissue (e.g., perpendicular to the plane of the leaf lamina, and at right angles to the mid-vein), the sample will need to be in a specific position for the process of ultramicrotomy.
-
1.
Once the resin has polymerised, if available, use an embedding capsule press (see Subheading 2.2, item 8) to remove the blocks from the moulds. Alternatively, very carefully cut down two opposing sides of the capsule with a single-edged razor blade and peel the two halves of the capsule apart to release each block.
-
2.
Using a simple dissecting microscope, locate (and mark if necessary) the position of the sample within each block (see Fig. 2a); it may help to draw precise guidelines on the block face for sawing along (see Note 29).
-
3.
Insert the block vertically, specimen up, into a small vice. Position the block to leave 3–4 mm of resin above the jaws of the vice. Close the vice (do not over-tighten it) and use a small hand saw (hacksaw) to remove the identified piece of resin containing the sample from the block (see Fig. 2b and c) (see Note 30).
-
4.
Remove any dust or sawing debris from both the excised piece of resin and the remaining block, hereafter called the base.
-
5.
Remove the base from the vice and position it vertically onto a piece of Blu-Tack pressure adhesive stuck atop a glass microscope slide or similar.
-
6.
Mix, according to the manufacturer’s instructions, approximately 0.5–1.0 mL of Araldite epoxy resin adhesive; place a small quantity of this adhesive onto the upper surface of the base.
-
7.
Using forceps, firmly insert the excised piece of resin containing the sample (longest edge to the base) into the Araldite adhesive. Ensure that the remounted piece is correctly orientated before building up the adhesive around all sides of the piece.
-
8.
Allow the epoxy resin adhesive to harden (usually 24 h), after which time the block is ready for sectioning (see Fig. 2d).
-
9.
Proceed to Subheading 3.4.
3.4 Ultramicrotomy and Staining
This is the process of obtaining ultra-thin sections (slices) of the sample for observation in the TEM. Because of the risk of physical damage to the sample from the initial specimen collection, it is often necessary to obtain and observe many grids of multiple sections so as to ascertain the optimum area of tissue preservation. Ultramicrotomy is a technique that requires much training and specialist equipment and as such must be approached with the help of experienced assistance. Many reference books are available that deal with the intricacies and problems of ultramicrotomy and staining (6, 14). A detailed method is beyond the scope of this chapter, but for general information, we include a brief summary of the technique as follows.
The block (remounted tissue sample) is first trimmed to remove excess resin from around the tissue. The aim is to produce a block face, approximately 2–3-mm square, from which semi-thick sections of about 0.5 μm can be taken using a glass or special diamond knife. Behind the cutting edge of the knife, there is a trough that is filled with water, upon which the sections float. Three or four of the thick sections are grouped together and transferred on a drop of water (using a wire loop of 3-mm diameter) to the approximate centre of a glass microscope slide. Move the slide to a hotplate (set to 80–90°C) to dry. Stain the sections with a suitable light microscopy stain, e.g., 1% (w/v) toluidine blue in 1% (w/v) sodium borate (14). To do this, cover the sections with a drop of staining solution and leave the slide on the hotplate for ∼2 min (do not allow the stain to dry out). Remove the slide from the hotplate and carefully rinse away the stain with distilled water. Dry the slide on the hotplate for ∼2 min before observation.
The sections are observed using a light microscope in order to determine the exact location within the sample that has been reached, and to locate the area to be used for EM. Once this has been identified, trim the block further to reduce the face to a trapezium shape similar in appearance to Fig. 2c (the longest edge should ideally be only 0.75–1.0 mm). Next, ultra-thin sections (60–90-nm thick) are cut for EM purposes. These are collected on the surface of a specimen support (grid) by picking up a clean grid with forceps, submerging it in the water trough, moving it below the sections, and lifting it up through the surface, thus picking up the sections on the grid. It is usual to collect three or four sections on any single grid.
Grids have a diameter of 3.05 mm, which is the limiting factor regarding the size of sample that can be observed by TEM. Grids are produced with many different architectures, e.g., square or hexagonal mesh, different mesh sizes (quoted as holes per inch), and metal compositions. The mesh size determines the support given to the sections and, along with the bar thickness, dictates the percentage transmittance of the grid; i.e., smaller mesh sizes (e.g., 300+ holes per inch) give greater section stability and support, but lower transmittance and therefore, potentially, a smaller viewable sample area. For routine TEM applications, a 300 square mesh copper grid may be chosen. Once the sections have been collected on a grid, they may be contrasted using heavy metal stains. For regular TEM observations, we routinely use the following staining protocol.
3.4.1 Primary Staining with Uranyl Acetate (see Notes 1 and 5)
If uranyl acetate has been used in the fixation process, this primary staining may be omitted.
In a fume cupboard, for each grid to be stained, pipette a 25–30 μL drop of filtered (0.2 μm) 2% aqueous uranyl acetate stain (see Subheading 2.5.3) onto a freshly exposed Parafilm surface; this is best done contained within a 9-cm diameter glass Petri dish. Using suitable forceps (see Subheading 2.2, item 2), submerge the grids in the stain, one grid per droplet. Then, cover the Petri dish by placing a box over it to exclude light and leave it for 15–20 min.
Using the forceps, remove each grid from the staining solution and wash by immersion, several times, in a substantial volume (10–20 mL) of distilled water contained in a 25-mL glass beaker. Repeat this process through at least three successive washes to remove all traces of stain. The washes must all be disposed of appropriately. Remove excess water from the grid using a small piece of filter paper and allow the grids to air-dry whilst still held by the forceps.
3.4.2 Secondary Staining with Reynolds Lead Citrate (12) (see Notes 1 and 12)
This process should be carried out in a nitrogen (carbon dioxide free) atmosphere; to this end, we utilise an in-house made Perspex box through which nitrogen gas is allowed to flow at 1–2 L per min (see Note 31).
From the filtered (0.2 μm) lead citrate stock (see Subheading 2.5.5), dispense one 25–30 μL drop of stain for each grid to be stained onto a freshly exposed Parafilm surface contained within a 9-cm glass Petri dish. Submerge the grids in this solution, one grid per droplet, and leave them for 2–3 min.
After staining, wash the grids as described in Subheading 3.4.1 by immersion in a substantial volume (10–20 mL) of freshly boiled distilled water (see Note 13).The first wash should contain one or two drops of 1 N sodium hydroxide to reduce the chance of lead carbonate precipitation. The grid should be passed through at least three successive boiled distilled water washes to remove all traces of stain. As with the primary stain, the washes must all be disposed of appropriately. Leave the grids to air-dry as before whilst still held by the forceps. It is advisable to store the stained grids in a labelled grid storage box (see Subheading 2.3, item 13) (see Note 32) prior to observation using the TEM.
3.5 Observation and Recording
Observation of the samples is carried out in the TEM, operated according to the manufacturer’s instructions. We routinely use an accelerating voltage of 80 kV, although this is just one of the parameters that may be adjusted in order to optimise results. We use an Olympus SIS Megaview III digital camera (Olympus Soft Imaging Solutions), which is controlled via dedicated iTEM software to capture and save images as required. Typical TEM images of chloroplasts in wild-type Arabidopsis cotyledons are shown (Fig. 3).
In order to enable meaningful comparisons between different genotypes, treatments, or conditions, it is essential to analyse a relatively large number of different organelles from multiple individuals (not least because of the nature of the sectioning process and the influence that it may have on what is observed). We recommend that at least three independent plants per genotype/treatment/condition are analysed, and that at least ten representative whole-organelle images per plant are selected for detailed analysis (the selection of representative images having been made on the basis of a careful overview of the whole section); i.e., a minimum of 30 independent, representative images per sample should be considered.
Images from the TEM can be used to derive quantitative data. For example, digital image files can be analysed using Adobe Photoshop or more specialised image analysis software. In Photoshop, the “Measure Tool” function can be used to obtain length and width measurements for each individual organelle. Such values can then be used to provide an indication of chloroplast shape (the length/width ratio; typically 3 in wild-type Arabidopsis), or to derive an approximate estimation of organellar cross-sectional area (using the following formula, which describes a perfect elliptical shape: π × 0.25 × length × width) (15, 16). The TEM images can also be used to derive other quantitative measures of development, such as the number of granal or stromal lamellae per granal stack.
4 Notes
-
1.
Sample processing for electron microscopy requires the handling of hazardous materials including chemical fixatives, solvents, embedding resins, and heavy metal solutions. Suitable personal protective equipment and the availability of approved containment facilities are essential. It is imperative to both consult and understand each material safety data sheet, and to ensure that appropriate approved codes of practice and local regulations for safe handling and waste disposal are strictly followed. Because of the significant hazards presented, persons using these materials must also be familiar with emergency procedures associated with accidental release or spillage. Ultimate disposal of many of these substances will require an approved chemical waste contractor.
-
2.
The proportions of formaldehyde and glutaraldehyde fixatives may be varied, as may the buffer vehicle (e.g., sodium cacodylate may be used in place of SP buffer). The fixation time, and the dehydration solvent (ethanol or acetone) and/or time may be adjusted and optimised for any particular specimen (17).
-
3.
Osmium tetroxide is very volatile and should always be used in a fume cupboard. It is also advisable to have a plentiful supply of milk powder available (or other suitable absorbent material) that can be used to absorb any spills. Glauert (9) further suggests that waste osmium may be converted to osmium dioxide (black) by mixing with an ethanol/water mixture or with ferrous sulphate. Other suitable methods of inactivation include mixing with vegetable oil or milk; Bozzola (18) recommends mixing in the ratio of two or three parts corn oil to one part osmium. All waste material must then be appropriately disposed of according to local regulations.
-
4.
Potassium ferricyanide (usually at 1.5% [w/v]) is an optional addition to the secondary fixative; its presence enhances membrane contrast (9).
-
5.
Uranyl acetate is particularly hazardous, especially in its powder form where inhalation presents a major hazard, as it is radioactive and very toxic, and may have cumulative effects. It must be handled according to local regulations, which should at the very least require containment to eliminate the risk of inhalation and ingestion. Uranyl acetate is also photolabile (9) and should, therefore, be stored in total darkness. It is quite slow to dissolve, so it is best prepared on the day before it is required.
-
6.
The original Spurr’s ultra low-viscosity resin (19) cannot now be purchased due to the withdrawal of one component, vinyl cyclohexene dioxide (VCD, ERL 4206). A modification to the original recipe is the replacement of this component with ERL 4221 (11).
-
7.
Spurr’s resin is particularly susceptible to moisture content. Therefore, to reduce the possibility of atmospheric moisture ingress, always keep component bottles closed when not in use and always cap the mixing vessel whilst stirring. Loaded specimen capsules should also be closed as quickly as possible. Water present in the final mix can lead to blocks of unsatisfactory consistency and prove problematic at the ultramicrotomy stage.
-
8.
Other epoxy resins are obtainable; for plant material, it is always advisable to use the lowest viscosity available.
-
9.
Some of the components used in these methods are very viscous and as such it is neither accurate nor practical to measure them by volume.
-
10.
We prefer plastic transfer pipettes because disposable glass pipettes may have fragments of broken glass that could be introduced into the resin and other solutions. If this occurs, there is a risk that when sectioning, a glass fragment embedded in the resin near the sample (which would be invisible) could damage the ultramicrotomy knife. Indeed, if using a diamond knife, this would destroy that area of the cutting edge.
-
11.
A decrease in the amount of DER 736 used in the mixture will give a harder block, while an increase will give a softer block. This effect may be used to determine the optimum hardness for any given sample.
-
12.
Lead citrate is readily precipitated (as lead carbonate) by the presence of carbon dioxide. It is best to use commercially purchased 1 N sodium hydroxide in the preparation of this stain, to ensure the correct pH of the final solution (13).
-
13.
To minimise the risk of lead carbonate precipitation, we prefer to use freshly boiled distilled water rather than stored distilled water that may have absorbed atmospheric carbon dioxide.
-
14.
Removing the top 30–50% of the organ is performed to assist penetration of the fixative. Single- or double-edged razors may be used. Double-edged blades are thinner than single-edged blades, and hence there is less potential for tissue damage at the point of cutting. Nonetheless, because of this possible damage, it is advisable to remove 0.5–1.0 mm of the tissue back from the immediate vicinity of the cut during the initial trimming of the block face at the ultramicrotomy stage.
-
15.
Processing containers should ideally be considered disposable, as they will become resin contaminated. Key points to consider when choosing a processing container are (A) resistance of the material to the various chemicals that will be used; (B) efficiency of containment of the chemicals – chemical vapour escape must be avoided; and (C) container volume. Consideration must be given to the efficacy of solution changes at each stage; the replacement solution may be diluted by residual solution from the previous stage. Therefore, to minimise this effect, volumes should not be reduced too much. Small screw-top containers are preferred, as they seal completely and present minimal risk of aerosol formation when opening (which can be a hazard with “snap-top” containers). Test any new container to ensure that it meets all requirements before use.
-
16.
A fine paintbrush is an excellent tool for manipulating the sample pieces; this is far less likely to cause the physical damage that might occur when using forceps.
-
17.
If available, it may help the infiltration of the fixative into the samples to expose them to a slight vacuum for the first 20–30 min of the primary fixation step (in fact, with some sample types this may be essential). This should not to be any more than 0.5 atmospheres (380 mmHg) (9). This should be applied and released intermittently with a frequency of 1–2 min per cycle (20).
-
18.
Because they contain hazardous aldehyde fixative, the sample tubes should ideally be contained within a sealed, labelled container.
-
19.
It is vital at all stages that the samples are not allowed to dry out; for this reason, solution exchanges should be completed one by one and as quickly as possible.
-
20.
The samples must be closely monitored. Leaf specimens have a tendency to “stick” to the sides of any container, and some may float until the 70% ethanol stage. This issue can be a particular problem when removing the previous solution at any exchange step, as the samples may adhere to the transfer pipette or tip. If this happens, a small amount of the next stage solution can be used to wash the adherent piece(s) carefully back into the processing tube.
-
21.
To avoid any possibility of crossover between samples (i.e., between different genotypes or conditions), always employ good laboratory practice and never use the same pipette to process different samples.
-
22.
All traces of phosphate must be removed from the samples at this stage. Residual phosphate may lead to uranyl phosphate precipitation, resulting in electron-dense deposits (21).
-
23.
Analytical grade ethanol should be opened immediately before use, and to limit atmospheric water absorption, the stock bottle should not be left open for any length of time. We advise the purchasing of bottles of the minimal volume available (usually 100 mL).
-
24.
1,2-Epoxypropane (PO) is extremely volatile; this poses two significant problems during pipetting. Firstly, any liquid drawn into a pipette starts to evaporate rapidly, filling the bulb with vapour which rapidly forces the liquid back out of the pipette. To prevent this, fill and drain the pipette two to three times (thus saturating the pipette bulb with vapour) before acquiring the required volume. The second problem can be the formation of condensation on the outside of the transfer pipette; this must never be allowed to enter either the sample tube or the stock bottle.
-
25.
Waste PO/resin mixtures may be left, uncapped, in the fume cupboard to allow the PO to evaporate. All of the waste resin may ultimately be combined and polymerised as per the sample blocks before appropriate disposal.
-
26.
Labels should be a paper band 3–4 mm deep with pencil writing (most inks will be removed by the resin), and this should be positioned around the circumference, half way down the capsule.
-
27.
To transfer each piece of leaf/cotyledon to an embedding capsule, modify a 7-mL transfer pipette by cutting off ∼0.5 cm from the tip. This gives a tube of larger diameter that should be sufficiently wide to draw each sample piece, individually, with just a small amount of the resin for transfer to the capsule.
-
28.
If the specimen is located to one side of the capsule, to simplify later remounting, lay the filled/closed embedding capsule on its side. The sample, visible through the base of the capsule, should slowly descend to the centre of the capsule (this should take no more than 2–3 min). When achieved, return the capsule to a vertical orientation. Alternatively, before closing the lid, a toothpick or similar may be used to position the sample very carefully.
-
29.
If deemed necessary because of size or visibility, a fine-tip marker pen may be used to draw a trapezium on the resin block face enclosing the sample. This may assist subsequent sawing/removal. Ensure that the longer of the two parallel edges is at 90° to the leaf stem (see Fig. 2).
-
30.
Sawing of resin blocks should be conducted in a fume cupboard to eliminate the risk of inhaling dust.
-
31.
If a nitrogen atmosphere chamber is not available, lead citrate staining may be conducted using a staining (Conway) dish comprising a central wax bed surrounded by an annulus “moat” of sodium hydroxide pellets in distilled water, and covered with a ground glass plate/lid.
-
32.
EM grids can be very susceptible to static charge. For this reason, it is advisable to store them in a dedicated grid storage box rather than in a Petri dish, where they may be readily attracted to the lid of the dish.
References
Whatley, J. M. (1978) A suggested cycle of plastid developmental interrelationships. New Phytol. 80, 489–502.
López-Juez, E., and Pyke, K. A. (2005) Plastids unleashed: their development and their integration in plant development. Int. J. Dev. Biol. 49, 557–577.
Kinsman, E. A., and Pyke, K. A. (1998) Bundle sheath cells and cell-specific plastid development in Arabidopsis leaves. Development 125, 1815–1822.
Schelbert, S., Aubry, S., Burla, B., Agne, B., Kessler, F., Krupinska, K., and Hörtensteiner, S. (2009) Pheophytin pheophorbide hydrolase (pheophytinase) is involved in chlorophyll breakdown during leaf senescence in Arabidopsis. Plant Cell 21, 767–785.
Kubis, S., Patel, R., Combe, J., Bédard, J., Kovacheva, S., Lilley, K., Biehl, A., Leister, D., Ríos, G., Koncz, C., and Jarvis, P. (2004) Functional specialization amongst the Arabidopsis Toc159 family of chloroplast protein import receptors. Plant Cell 16, 2059–2077.
Kuo, J. (2007) Electron Microscopy: Methods and Protocols. Methods in Molecular Biology, Vol. 369. Humana Press, Totowa, NJ, USA.
Pfeiffer, S., and Krupinska, K. (2005) Chloroplast ultrastructure in leaves of Urtica dioica L. analyzed after high-pressure freezing and freeze-substitution and compared with conventional fixation followed by room temperature dehydration. Microsc. Res. Tech. 68, 368–376.
Hayat, M. A. (1981) Fixation for Electron Microscopy. Academic Press, New York, USA.
Glauert, A. M., and Lewis, P. R. (1998) Biological Specimen Preparation for Transmission Electron Microscopy. Practical Methods in Electron Microscopy, Vol. 17. Portland Press Ltd., London, UK.
Li, H., Culligan, K., Dixon, R. A., and Chory, J. (1995) CUE1: a mesophyll cell-specific positive regulator of light-controlled gene expression in Arabidopsis. Plant Cell 7, 1599–1610.
Ellis, E. A. (2006) Solutions to the problem of substitution of ERL 4221 for vinyl cyclohexene dioxide in Spurr low viscosity embedding formulations. Microscopy Today 14, 32–33.
Reynolds, E. S. (1963) The use of lead citrate at high pH as an electron-opaque stain in electron microscopy. J. Cell Biol. 17, 208–212.
Ellis, E. A. (2007) Poststaining grids for transmission electron microscopy: conventional and alternative protocols. In, Electron Microscopy: Methods and Protocols (Kuo, J., ed.) Humana Press, Totowa, NJ, USA, pp. 97–106.
Hunter, E. (1993) Practical Electron Microscopy: A Beginner’s Illustrated Guide. Cambridge University Press, Cambridge, UK.
Kovacheva, S., Bédard, J., Patel, R., Dudley, P., Twell, D., Ríos, G., Koncz, C., and Jarvis, P. (2005) In vivo studies on the roles of Tic110, Tic40 and Hsp93 during chloroplast protein import. Plant J. 41, 412–428.
Aronsson, H., Boij, P., Patel, R., Wardle, A., Töpel, M., and Jarvis, P. (2007) Toc64/OEP64 is not essential for the efficient import of proteins into chloroplasts in Arabidopsis thaliana. Plant J. 52, 53–68.
Hall, J. L., and Hawes, C. (1991) Electron Microscopy of Plant Cells. Academic Press, London, UK.
Bozzola, J. J. (2007) Conventional specimen preparation techniques for transmission electron microscopy of cultured cells. In, Electron Microscopy: Methods and Protocols (Kuo, J., ed.) Humana Press, Totowa, NJ, USA, pp. 1–18.
Spurr, A. R. (1969) A low-viscosity epoxy resin embedding medium for electron microscopy. J. Ultrastruct. Res. 26, 31–43.
Kuo, J. (2007) Processing plant tissues for ultrastructural study. In, Electron Microscopy: Methods and Protocols (Kuo, J., ed.) Humana Press, Totowa, NJ, USA, pp. 35–45.
Louw, J., Williams, K., Harper, I. S., and Walfe-Coote, S. A. (1990) Electron dense artefactual deposits in tissue sections: the role of ethanol, uranyl acetate and phosphate buffer. Stain Technol. 65, 243–250.
Acknowledgments
The authors wish to acknowledge Ms. Natalie Allcock of the Core Biotechnology Services Electron Microscopy Laboratory, University of Leicester, for both technical support and comments on the manuscript.
Author information
Authors and Affiliations
Corresponding author
Editor information
Editors and Affiliations
Rights and permissions
Copyright information
© 2011 Springer Science+Business Media, LLC
About this protocol
Cite this protocol
Hyman, S., Jarvis, R.P. (2011). Studying Arabidopsis Chloroplast Structural Organisation Using Transmission Electron Microscopy. In: Jarvis, R. (eds) Chloroplast Research in Arabidopsis. Methods in Molecular Biology, vol 774. Humana Press. https://doi.org/10.1007/978-1-61779-234-2_8
Download citation
DOI: https://doi.org/10.1007/978-1-61779-234-2_8
Published:
Publisher Name: Humana Press
Print ISBN: 978-1-61779-233-5
Online ISBN: 978-1-61779-234-2
eBook Packages: Springer Protocols