sample preparation of sem for plant smaples

ISSN:1369 7021 © Elsevier Ltd 2009VOLUME 12 – ELECTRON MICROSCOPY SPECIAL ISSUE32

Sample preparation for SEM of plant surfaces

Scanning electron microscopy (SEM) is an ideal technique for

examining plant surfaces at high resolution. Plant tissues must

be preserved by dehydration for observation in an electron

microscope because the coating system and the microscopes

operate under high vacuum and most specimens cannot withstand

water removal by the vacuum system without distortion1.

In order to examine the native structure of the sample, some

microscopes are designed to image frozen hydrated samples and

Plant tissues must be dehydrated for observation in most electron microscopes. Although a number of sample processing techniques have been developed for preserving plant tissues in their original form and structure, none of them are guaranteed artefact-free. The current paper reviews common scanning electron microscopy techniques and the sample preparation methods employed for visualisation of leaves under specific types of electron microscopes. Common artefacts introduced by specific techniques on different leaf types are discussed. Comparative examples are depicted from our lab using similar techniques; the pros and cons for specific techniques are discussed. New promising techniques and microscopes, which can alleviate some of the problems encountered in conventional methods of leaf sample processing and visualisation, are also discussed. It is concluded that the choice of technique for a specific leaf sample is dictated by the surface features that need to be preserved (such as trichomes, epidermal cells or wax microstructure), the resolution to be achieved, availability of the appropriate processing equipment and the technical capabilities of the available electron microscope.

A.K. Pathana*, J. Bondb and R.E. Gaskina

aPlant Protection Chemistry NZ, PO Box 6282, Rotorua, New ZealandbSCION, Private Bag 3020, Rotorua, New Zealand

*Email: [email protected]

MTElectronMicro_p32_43.indd 32 08/07/2010 11:52:16

Open access under CC BY-NC-ND license.

mailto:[email protected]

http://creativecommons.org/licenses/by-nc-nd/3.0/

Sample preparation for SEM of plant surfaces REVIEW

VOLUME 12 – ELECTRON MICROSCOPY SPECIAL ISSUE 33

more recently environmental SEM microscopes have been developed

which can image the sample in their native-hydrated state. These

microscopes are specialised equipments and may not be available

in many labs. Hence, sample preparation by dehydration is still an

important consideration for observation in conventional microscopes.

For samples that necessitate dehydration, many techniques other

than just air-drying have been developed to remove water from the

sample, all aiming at minimal distortion of the cell and maximal

preservation of the original form and structure. These techniques

include freeze-drying, critical point drying, and various types of

chemical fixation treatments prior to dehydration of samples. However,

acceptable methods offer less than ideal preservation for some

plant species and may be inconsistent. The inconsistency is largely

due to diversity in tissue types, form, structure and composition of

plants. Inconsistencies also arise from variation in individual skills and

equipment used across different labs. Hence, new/modified techniques

are continually being tested and developed for the preparation of

specific plant tissues for visualisation under electron microscopes.

The paper reviews common techniques/methods used in the past

for leaf sample preparation for scanning electron microscopy. Selected

examples from our own work on common plant species (monocots and

dicots) of interest in pesticide research are presented and compared

with those from previous studies that have used similar techniques

for electron microscopy of plant tissues. Emphasis has been given to a

simple, but robust leaf sample preparation technique (simple air-drying),

which has proved highly effective for visualisation of plant waxes under

a field emission scanning electron microscope (FESEM) at low kV.

Approaches for sample preparation and visualisationSamples can be visualised in their native-hydrated state without

pre-treatment, frozen hydrated state or after removing liquids from

the samples using a variety of techniques. The choice of technique

will depend on the sample, the equipment available and the surface

features and structures that need to be visualised.

Hydrated samplesRapid observations of fresh hydrated samples can be made by

using an environmental scanning electron microscope (ESEM). The

technique has the potential to provide excellent low magnification

images of plant surfaces in their native-hydrated state. In addition, it

allows the flexibility to alter stage temperature and vapour pressure

in the specimen chamber. For example, leaf tissues can be examined

at high humidity in the chamber and minimise sample dehydration

during the imaging process. This technique can also be effectively used

to perform ‘dynamic’ experiments in wet mode to examine biological

events in developmental processes such as fungal growth on leaf

surfaces.

A FEI Quanta ESEM† (FEI Company, USA) was used at an

accelerating voltage of 10–20 kV, a stage temperature of 2 °C and a

chamber pressure of 6 Torr to visualise unprocessed chenopodium and

pea leaf surface in their native-hydrated state (Fig. 1a–d). Although the

‘true-to-life’ low magnification images of chenopodium leaves were

† Equipment used at Research Centre for Surface and Materials Science, University of Auckland, New Zealand.

Fig. 1 Leaf surfaces from unprepared and uncoated specimen visualised under an environmental scanning electron microscope: (a) chenopodium leaf surface showing intact epidermal cells and salt glands; (b) chenopodium salt gland at high magnification, note that waxes are not visible using this technique; (c) pea leaf surface showing intact epidermal cells, but waxes are not clearly visible; (d) epidermal cell collapse in pea leaf surface at high magnification. Waxes are not clearly visible.

(b)(a)

(c) (d)


REVIEW Sample preparation for SEM of plant surfaces

VOLUME 12 – ELECTRON MICROSCOPY SPECIAL ISSUE34

successfully obtained, the wax microstructure on the salt gland could

not be seen at all (compare Fig. 1 and Fig. 9) using this technique.

Similarly, the wax microstructure on pea leaf surface was not as clearly

visible as that from carefully air-dried samples (compare Fig. 1 and

Fig. 6). The sample also collapsed during imaging of the surface at high

magnification (5000×, Fig. 1d).

Despite having some limitations, the technique may allow

examination of some samples that could never be viewed in a

conventional SEM in their true-to-life form. This technique is increasingly

being used to study trichomes and glandular morphology and

function2-5. It is also being used to study elemental distribution in fresh

leaf samples using an appropriate energy-dispersive X-ray spectroscopy

(EDS) detector. Silicon accumulation in leaf tissues of sorghum, rice,

bamboo and orchard grass has been successfully studied using this

technique6-9. However, it is anticipated that the EDS scanning times

could be restricted to shorter lengths since the samples are relatively

delicate and unstable as compared to the dried samples that are robust

and suited for long time EDS examinations in a conventional SEM.

ESEM also provides a useful tool in taxonomic studies to classify

insects even at the species level10. The mechanism of leaf penetration

by fungi is not completely understood and may vary between

pathogens11. Studies with an ESEM may be useful to decipher the

fungal infection processes better. The technique can also be used to

study water droplet interactions on foliate surfaces. Cheng et al.12

successfully observed water condensation and evaporation processes on

lotus leaf surface using an ESEM and developed a geometric model for

liquid drops on rough surfaces.

Inherent limitations associated with ESEM include reduced field of

view and depth of focus, and in many cases, reduced resolution and

stability at high magnification since the sample is viewed without

coating with a heavy metal such as gold or chromium. Some samples

may move/change form during examination and necessitate rapid or

repeat image capture for a satisfactory micrograph. This equipment can

be considered as a SEM with added degrees of difficulty; however, the

art of imaging using an ESEM may be perfected for specific samples

using the right combination of accelerating voltage, stage temperature,

vapour pressure and working distance.

As an alternative to ESEM, a conventional SEM can be used to

view hydrated samples using special techniques such as QuantomiX

WetSEM™ technology that comprises a vacuum-tight capsule bounded

by a unique, electron-transparent and pressure-resistant membrane

(www.quantomix.com). This system allows visualisation of a wide

variety of fully hydrated samples in a conventional SEM by isolating the

sample from the vacuum while allowing penetration and reflection of

a scanning electron beam. A major disadvantage of this technique is its

relatively high cost due to the fact that a capsule can normally be used

only once for each sample.

Hydrated samples can also be visualised in a conventional SEM

if the liquids in the samples are substituted by glycerol13. Glycerol

evaporates very slowly under vacuum as compared to water, thereby

allowing the sample visualisation for short time in a conventional SEM

without causing malfunction of the vacuum system. An advantage of

the glycerol substitution method proposed by these authors is that the

leaf sample can be infiltrated from the underside by mounting them on

a piece of fabric soaked with glycerol, thus leaving the upper surface

untouched for visualisation. However, hydrophobic samples may prove

difficult to infiltrate with glycerol and develop shrinkage artefacts13.

This may not be a problem if the lower leaf surface is hydrophilic

and allows glycerol infiltration with ease. One good example of such

species is ryegrass, with the lower leaf surface much more wettable

than the upper leaf surface. A disadvantage of this technique is that the

longer visualisation times (greater than 30 min) may result in glycerol

accumulation in the oil of the vacuum pumps causing malfunction

of the equipment. In addition, satisfactory EDS analysis may not be

possible due to the scanning time restriction to ca. 30 min and the

possibility of changes in elemental form and distribution introduced by

the glycerol substitution. Although this technique has not been tested

in our lab due to intolerance of the FESEM to any kind of wet sample, it

was successfully used by Wagner et al.14 and Koch et al.15 to visualise

surface features of a range of plant species using a conventional SEM.

Recently a new TM-1000 tabletop microscope (Hitachi High-

Technologies Corporation, Japan)16 has been introduced that is a

relatively low-cost, portable, easy-to-use ESEM with a magnification

range of 20–10,000×. In addition, the TM-1000 detector is also

capable of showing contrast arising from differences in atomic number.

This instrument can be portable and carried to field sites for in situ

examination of plants, insects and microbes.

Frozen hydrated samplesFresh samples may be frozen (cryofixation) in their native-hydrated

state, coated and visualised on a cold-stage (cryo-chamber) attached

to the SEM. This technique, commonly called low temperature scanning

electron microscopy (LTSEM), has been widely used in the past for

examining plant surfaces. It is quick and is best suited to samples with

higher water content—samples that are usually very difficult to process

by conventional methods. This method is useful where preservation

of the natural ‘life-like’ morphology of cells and tissues is desired17.

Cryofixation is rapid and immobilises processes at a much faster rate

than chemical fixation18. This is a big advantage if dynamic biological

processes such as fungal spore discharge19 or fungal infection of a leaf20

need to be captured step-by-step in a ‘suspended animation’ form.

Another potential application of LTSEM is to trace elements on

plant surfaces or in freeze-fractures using EDS21-23. Some of these

elements may otherwise be lost to physiological processes (e.g.

translocation, leaching or biodegradation) or solubilised/moved

by solvents or chemicals if samples are processed by conventional

methods. Rapid freezing helps to retain them in their original form and

location.


http://www.quantomix.com



A Philips SEM 505‡ coupled to a Hexland cryopreparation system

at a temperature of −140 °C to −120 °C was used at an accelerating

voltage of 4 kV to observe bean and wheat surfaces24. The samples

were attached to the stub and frozen by sample-holder contact with

the pre-chamber stage (−170 °C) in an atmosphere of dry nitrogen.

The samples were first viewed at low kV and ice contamination (if any)

was removed by raising the stage temperature to −60 °C prior to

coating with a gold layer of ca. 20-nm thickness. Excellent artefact-free

images of the leaf surface were obtained for both species with minimal

surface distortion as evident from intact epidermal cells and stomata

on bean and wheat leaves (Fig. 2a and b).

LTSEM can also be effectively used to observe the form of pesticide

deposit on plant surfaces25, since the deposits are cryo-fixed in their

original form and location without being exposed to chemical fixatives,

solvents or dehydrating forces as in conventional sample preparation

methods. The form and distribution of deposits remains unchanged

during visualisation as opposed to that under an ESEM in which it

may change due to introduction of additional moisture in the ‘wet’

mode. Using LTSEM, foliar deposits of the herbicide glyphosate with

and without adjuvants (spreading/penetrating and non-spreading/

non-penetrating) were observed on wheat leaf surfaces. The semi-

crystalline herbicide deposits and their modification by the adjuvants

on wheat leaf surface were effectively photographed in their original

form (Fig. 2c–f24).

LTSEM may also be used to examine internal structures by freeze-

fracture. However, the possibility of damage due to internal ice-crystal

growth needs to be considered. Rapid freezing (at several 1000 K/s)

of samples is required to minimise ice-crystal formation, or in ideal

Fig. 2. Leaf surfaces ((a) bean and (b) wheat) examined using a low-temperature scanning electron microscope (LTSEM). (c–f) Foliar deposits of herbicide glyphosate on wheat (with and without adjuvant) examined using a LTSEM in frozen hydrated state 1 h after application of the herbicide. (c) The herbicide deposit (without adjuvant) has formed a dense semi-crystalline film on top of epicuticular wax crystals (edge marked by arrows) and has failed to wet stomatal depressions; (d) higher magnification of the deposit edge of (c) showing irregular shaped semi-crystalline aggregates of herbicide (edge marked by arrows) formed on epidermal surface; (e) edge of herbicide deposit (marked by arrows) containing a spreading and penetrating adjuvant. Deposit is difficult to detect due to extensive spread and penetration caused by the adjuvant; (f) edge of herbicide deposit (marked by arrows) containing non-spreading and non-penetrating adjuvant. Deposit is clearly visible as an amorphous film lying on top of the epicuticular wax crystals.

‡ Equipment used at Long Ashton Research Station, Bristol, U.K.

(b)(a)

(c) (d)

(e) (f)




conditions, to reduce water to a glassy state (vitreous ice). The faster

the cooling, the smaller the ice crystals will be. Crystal sizes of less

than 10 nm will do little damage to the samples. The rate of freezing

will also depend on the tissue thickness and composition. A number

of techniques can be utilised for rapid freezing, including plunge

freezing in liquid nitrogen (standard method), spray freezing with

propane, propane jet freezing, ‘slam’ freezing against a liquid helium or

nitrogen-cooled polished metal block, high pressure freezing (>2100

bar), etc.26,27. However, the depth of the specimen that is free from ice

crystal damage is normally limited to the outermost layers, typically

15–20 μm depending on the tissue type28.

LTSEM has been extensively used in the past and the subject has

been reviewed in detail by various authors17,29. It is well established that

LTSEM provides superior images but the technique is also not entirely

artefact-free. Artefacts in LTSEM may originate during cryofixation,

etching, freeze-fracturing, coating, specimen transfer and electron beam

irradiation17. Artefacts may also be introduced when frozen hydrated

samples have to be partially etched to remove ice contamination from

fracture faces. The frozen specimens are very beam sensitive and beam

damage may cause cracking of the specimen surface30. In order to

utilise this technique to its full potential, good operational skills are

required for artefact-free imaging of hydrated samples. In addition, the

need for a specialised cryo-chamber limits the use of this technique to

the few labs that can afford the additional cost involved.

Dried/dehydrated samplesStandard SEM procedures for biological samples involve chemical

fixation, drying/dehydration, mounting on a stub and coating with a

metal (e.g. chromium, gold, platinum, etc.) for examination under a

conventional SEM, often referred as ambient temperature scanning

electron microscopy (ATSEM). The fixation, drying/dehydrating steps

need to be done as carefully as possible to reduce shrinkage while

ensuring preservation of cell structures as close to the natural state as

possible.

Fig. 3. Leaf surfaces from samples prepared using the CPD technique. (a) Bean leaf surface; (b) bean stomata; (c) chenopodium leaf surface; (d) chenopodium stomata; (e) broccoli leaf surface; (f) broccoli stomata. Note that some shrinkage of the epidermal cells is evident, especially around stomata. Waxes on chenopodium and broccoli appear to have been solubilised.

(b)(a)

(c) (d)

(e) (f)




There are several methods for drying/dehydrating leaf samples

for ATSEM, each having its own advantages and disadvantages. The

common drying techniques used in the past are (i) critical point

drying (CPD) (ii) freeze-drying (lyophilising) after prefreezing the

samples in liquid nitrogen-cooled liquid propane or Freon 22, and

then plunge freezing in liquid nitrogen31 and (iii) chemical fixation

in glutaraldehyde/osmium tetroxide before carrying out standard

dehydration in an organic solvent followed by CPD. To achieve an

acceptable preservation of plant tissues, these techniques have been

tried (with some variations) in different labs, including ours, with mixed

success32-37.

In our lab, the dehydrated samples were mounted on aluminium

stubs using aqueous conductive silver, and chromium coated (once

or twice) using an Emitech K575X peltier cooled turbo sputter coater

(Emitech Ltd., U.K.) prior to visualisation under a JEOL JSM-6700F field

emission scanning electron microscope (JEOL Ltd., Japan). The FESEM

is equipped with two secondary electron detectors: LEI (lower detector)

and SEI (upper detector). The SEI detector is higher in the column

and sees fewer shadows. Less charging is detected, and since it can be

used at a shorter working distance, the resolution is greater than with

the LEI. All surface wax images (at high magnifications) were taken

using the SEI detector since it gives comparatively high resolution. The

conditions used were an accelerating voltage 3–10 kV; illuminating

current 2.6 nA with a working distance 8–15 mm.

Critical point drying

Initially introduced by Anderson38 more than half-a-century ago, CPD

is the most commonly used dehydrating method for biological sample

preparation. This procedure removes liquids from the specimen and

avoids surface tension effects (drying artefacts) by never allowing a

liquid/gas interface to develop. The transition from liquid to gas at the

critical point takes place without an interface because the densities of

liquid and gas are equal at this point.

We used the standard CPD protocol for processing our samples.

After fixation with 2.5% glutaraldehyde in 0.2 M cacodylate and 2%

buffered osmium tetroxide, the samples were dehydrated through a

graded series of ethanol (10%, 20%, 30%, 50% and 70%—once for

10 min at each step), and then immersed in 100% acetone twice for

30 min each. The tissues were then transferred to an Emitech K850

critical point dryer (Emitech Ltd., U.K.) using liquefied carbon dioxide as

transitional fluid. We found that CPD gave an acceptable preservation

of bean, chenopodium and broccoli leaf surface (Fig. 3a, c and e,

respectively) but the organic solvents stripped-off epicuticular waxes

from chenopodium and broccoli (Fig. 3d and f, respectively). Some

shrinkage on leaf surface was also evident around stomata of bean

and chenopodium (Fig. 3b and d, respectively). Shrinking artefacts may

be introduced in the samples while permuting the specimens from

one solution to the next or into the CPD. To avoid these artefacts, it

is essential that the specimens are completely wet during the whole

preparation process. Although shrinking surface artefacts were observed

on some CPD processed samples, the internal structures (mesophyll

cells, etc.) were highly preserved in bean, chenopodium and broccoli

(Fig. 4a–c). A similar level of preservation of mesophyll cells was not

observed in a freeze-dried fracture of broccoli (Fig. 4d).

Bray et al.35 used CPD, Peldri II and hexamethyldisilazane (HMDS)

to determine a method that gave the best preservation of leaf tissues.

Fig. 4. Fractures from samples preservation using the CPD technique. (a) Bean; (b) chenopodium; (c) broccoli; (d) fracture of broccoli leaf surface preserved with the freeze-drying technique (liquid nitrogen) for comparison. Note the excellent level of preservation of mesophyll cells achieved using the CPD technique.

(b)(a)

(c) (d)




Peldri II caused complete extraction of leaf epicuticular wax, while CPD

and HMDS showed minimal extraction compared with that of samples

air-dried directly from acetone. They also found that while all the three

methods showed signs of shrinkage, CPD provided relatively better

quality of tissue preservation.

CPD is the method of choice (with careful use) if fractures

and/or non-waxy epidermal surfaces (e.g. bean leaf surface) are

to be examined in an ATSEM. It is distinctively advantageous for

specific applications and has been used widely for biological sample

preparation. However, the technique necessitates the use of a

specific apparatus and the sample throughput is limited39. It may not

completely remove water from some tissues and may cause some

bulk shrinkage or generate violent bubbling40,41. The technique also

necessitates the use of organic solvents such as acetone that may

damage leaf epicuticular wax structures42. These structures are an

important part of the leaf surface micro-morphology and need to be

adequately preserved for scanning electron microscopy.

Freeze-drying

The first step in freeze-drying technique is to rapidly freeze the sample

to avoid ice-crystal formation. Rapidity of freezing is probably the most

influential factor on the final preservation quality of biological samples.

Ideally, samples should be directly plunged into liquid nitrogen (−196 °C),

but liquid nitrogen forms a gaseous/insulating layer (Leidenfrost effect)

thus losing contact with the sample. To avoid the Leidenfrost effect,

samples can be plunged in slush nitrogen (−210 °C) prepared by placing

a beaker filled with liquid nitrogen in a desiccator under vacuum.

Alternatively, rapid freezing of samples can be achieved by plunging

in nitrogen-cooled Freon 22 (−150 °C) or liquid propane (−178 °C),

subsequently freezing in liquid nitrogen and then freeze-drying

Fig. 5 Leaf surfaces from samples prepared using freeze-drying technique; (a–d) rapid freezing achieved by plunging in liquid nitrogen-cooled liquid Freon 22. (a) Some preservation of bean epidermal cells; (b) wheat waxes relatively more preserved as compared to those of broccoli; (c) wax dissolution apparent in waxy cabbage leaf; (d) waxes solubilised around broccoli stomata; (e) rapid freezing of bean leaf sample achieved using liquid nitrogen slush. All samples were freeze-dried in a basic freeze-drying system using dry-ice cake under vacuum.

(b)(a)

(c) (d)

(e)




under controlled conditions. Some labs recommend introduction

of a cryo-protectant (usually 20–30% glycerol) into the tissues

to minimise ice crystal size. However, this technique necessitates

pre-fixation of samples in glutaraldehyde which may not be desirable

if chemical fixatives need to be avoided. In addition, cryo-protectants

may not completely sublime away with water during the freeze-drying

step.

Freeze-drying necessitates availability of a good (turbo pumped)

vacuum system, an effective cold trap (−80 °C to −100 °C) for

sublimed water and liquid N2-based cooling. Sophisticated freeze-

drying equipment is available (e.g. EMITECH K750 and K775; Emitech

Ltd., U.K.) that provides adequate flexibility to manipulate conditions

for specific sample types. For good sample preservation, both freezing

(needs to be rapid) and freeze-drying processes need to be optimised

for specific samples.

If a suitable freeze-dryer is not accessible, a basic freeze-drying

system can be developed in-house. In our study, cryo-fixed (frozen

in liquid nitrogen/slush or liquid nitrogen-cooled Freon 22) samples

were freeze-dried using an in-house simple freeze-drier. The sample

was placed on a liquid N2-cooled brass metal holder under vacuum

with an effective cold trap and allowed to dry overnight. We placed

a brass metal holder on a cake of dry ice under vacuum to maintain

the drying temperature below −60 °C. The cake gradually decreases

in size over time while the brass metal holder gradually sinks-in and

remains in complete contact with dry ice all the time, thus providing

a constant low temperature for freeze-drying. We found acceptable

preservation of bean (non-waxy) and wheat (waxy) samples using this

technique after freezing samples in liquid nitrogen-cooled liquid Freon

22 (Fig. 5a and b). However, it did dissolve waxes of other species such

as cabbage and broccoli (Fig. 5c and d). In theory, freezing samples

Fig. 6. Leaf samples prepared by chemical fixation with glutaraldehyde and osmium tetroxide followed by dehydration in ethanol and air-drying (left) and air-drying (right) techniques. (a and b) Barnyardgrass; (c and d) pea; (e and f) chenopodium. Note the relatively better preservation of leaf surfaces by simple air-drying as opposed to severe distortion by chemical fixation followed by dehydration in ethanol and then air-drying.

(b)(a)

(c) (d)

(e) (f)




in liquid nitrogen slush should provide good freezing and acceptable

preservation of samples after freeze-drying without the need to use

liquid nitrogen-cooled liquid Freon 22. However, adequate preservation

of the majority of bean leaf surface was not achieved by using this

technique in our experiments (Fig. 5e).

In our view, the techniques described above can be further

improved and perfected for specific samples using liquid Freon 22/

liquid propane (for many non-waxy plant species) or liquid nitrogen

slush (for waxy plant species); freeze-drying using a dry ice cake may

provide adequate low temperature conditions required for the drying

process if sophisticated freeze-drying equipments are not accessible. A

limitation to this technique is that the use of liquid Freon 22 and liquid

propane could be restricted in many labs since the former is a powerful

greenhouse gas and the latter is highly flammable.

Chemical fixation

Chemical fixation involves soaking samples for various periods

of time (depending on their thickness and composition), usually

in glutaraldehyde and/or osmium tetroxide. The samples may be

subsequently dehydrated in a graded series of ethanol32, and then

dried by CPD or subjected to air-drying. Alternatively, the sample may

be transferred to tetramethylsilane (TMS) for 10–20 min before air-

drying34.

We fixed bean samples in 2.5% glutaraldehyde and 2% osmium

tetroxide followed by dehydration in a graded series of ethanol. The

samples were then dried by two different methods—CPD and simple air-

drying (in a desiccator under vacuum). CPD has been already described

in an earlier section. Although, acceptable preservation was achieved

by the CPD technique post-fixation with glutaraldehyde and osmium

tetroxide, epidermal cells appeared to be shrunken, especially around

stomata of bean and chenopodium (Fig. 3b and d). This shrinkage

of tissue probably occurred during dehydration rather than drying43.

Hardy et al.44 also failed to achieve acceptable surface preservation for

Dactylis glomerata and Elymus canadensis leaf samples using CPD and

TMS techniques after fixation in glutaraldehyde/acrolein mixture.

Simple air-drying after fixation in glutaraldehyde and osmium

tetroxide caused substantial alterations of the leaf surfaces of

barnyardgrass, pea and chenopodium species (Fig. 6a, c and e). This

was despite the expectation that fixing in glutaraldehyde and osmium

should strengthen the elastic cell walls by cross-linking polymers and

thus resist cell collapse due to dehydrating forces. Since comparatively

less surface distortion was achieved by CPD (post-fixation with

glutaraldehyde and osmium tetroxide, Fig. 3a–f) as well as simple air-

drying without fixation (Fig. 6b, d and f), it is clear that the fixatives

negatively influenced the drying step to cause substantial damage. Bray

et al.35 concluded that post-fixation with osmium tetroxide generally

resulted in poorer specimen preservation than using Karnovsky’s

fixative (mixture of glutaraldehyde and formaldehyde), especially for

dicotyledonous species such as Coleus blumei using the CPD method.

The authors suggested that the increased distortion of plant cell

surface structure following post-fixation with osmium tetroxide might

be due to (i) a build up of osmium molecules that could ultimately

inhibit infiltration of dehydrating agents (e.g. ethanol, acetone,

Freon 113) and transitional agents (Freon 13, Freon 23, carbon dioxide)

Fig. 7. Bean leaf specimens prepared by (a) using methanol as a fixative followed by air-drying; (b) freeze-drying post-fixation with liquid nitrogen-cooled methanol; (c) fixation in glutaraldehyde and osmium tetroxide followed by dehydration in ethanol and air-drying. Note the different degrees and types of leaf surface distortions in (a–c) as compared to (d) where imaging was done using LTSEM.

(a) (b)

(c) (d)




in the CPD technique and (ii) the drying solvents (e.g. HMDS, Peldri

II, TMS or dimethoxypropane—DMP) used as an alternative to the

CPD technique45-47. It is recommended that leaf samples should not

be fixed with glutaraldehyde and/or osmium tetroxide if post-fixation

drying is to be achieved by simple air-drying or with organic solvents.

Methanol was proposed as an alternative fixative/dehydrant that

could be used prior to CPD for preserving plant surfaces37. The use of

methanol instead of glutaraldehyde or osmium tetroxide was suggested

since it instantly fixes the elastically extended cell walls and can rapidly

penetrate inside plant cuticles and cell walls. The authors suggested that

this method resulted in improved fixation of cell wall dimension and is

deemed to be the most suitable for preserving plant epidermal surfaces.

They also achieved superior preservation of Silvina auriculata and

Verbascum arcturus trichomes as compared to conventional treatments.

We tested a variation of this technique by (i) immersing bean

samples in methanol for 20–40 s followed by simple air-drying (Fig. 7a)

or by (ii) immersing in liquid nitrogen-cooled methanol for 20–40 s,

re-immersing in liquid nitrogen followed by freeze-drying (Fig. 7b).

The techniques provided preservation of epidermal cell surface in

some areas and collapse in others, but the latter technique (Fig. 7b)

appeared to be relatively better in preservation quality. Although

the quality of preservation was far from ideal, it was still better than

that obtained from using glutaraldehyde and osmium tetroxide as

fixatives (Fig. 7c).

We expect that the methanol fixation of plant tissues to have

some potential for fine tuning for CPD, freeze-drying or simple air-

drying techniques and to have an application in preserving specific

plant tissues. Use of methanol as a fixative in these processes is also

Fig. 8. Wax micro-morphology of specimens processed by simple-air drying and observed under a FESEM. (a) Barnyardgrass leaf; (b) grape berry; (c) broccoli leaf; (d) cabbage leaf; (e) stephanotis leaf; (f) hedera leaf; (g) barley leaf; (h) pea leaf. Note the high resolution and brightness achieved by using this technique with minimal wax disruption.

(b)(a)

(c) (d)

(c) (d)

(e) (f)




desirable for waxy species since it is relatively less damaging to plant

waxes as compared to ethanol or acetone.

Simple air-drying without pre-treatment

In order to avoid problems in sample preparation described above and

due to lack of access to a suitable LTSEM, we tried the simplest sample

preparation procedure—air-drying without pre-treatment. Samples

were slowly allowed to dry at room temperature in a desiccator for

12–24 h (depending on the species used and inherent turgidity in their

leaf tissues) with or without vacuum. Some shrinkage of the epidermal

cells was observed as expected (Fig. 6b, d and f). However, wax

microstructure could be successfully examined at high resolution (up to

50000×, Fig. 8a–h).

The wax images were taken using a field emission scanning electron

microscope that allows high-resolution imaging at low electron dose.

Using this technique, chenopodium leaf surface features could also be

examined at high resolution (Fig. 9a–d). The similarity in wax micro-

morphology of chenopodium leaf and salt gland is quite evident from

these micrographs (Fig. 9c and d). To our knowledge, this is the first

published image of wax microstructure on chenopodium glands at high

resolution.

The simple air-drying technique in combination with a FESEM

worked best for high-resolution imaging of wax microstructure of a

range of plant species. The air-dried samples (and the waxes) were quite

stable even at high electron dose and accelerating voltages, making it a

technique ideally suited for high-resolution imaging of plant waxes.

Summary and conclusionsThe dehydration of plant tissues for scanning electron microscopy poses

distinctive challenges. A range of sample preparation and visualisation

techniques can be employed to overcome difficulties arising from plant

tissue characteristics, but none of them are guaranteed artefact-free.

Artefact-free preservation of plant specimens is difficult to achieve due

to inherent plant tissue characteristics such as hydrophobic cuticles

and thick cell walls that impede penetration of aqueous fixatives.

Additionally, the presence of a large central vacuole that can dilute

fixative/buffer concentrations or collapse after water is removed by

dehydration, and low protein content of cells that impart reduced cross-

linking with glutaraldehyde, etc. impose further restrictions.

Variations in plant specimen preservation also arise due to

equipment capabilities as well as individual skills and expertise. Fresh,

uncoated material can be examined directly in an ESEM, but there

may be constraints over resolution and magnification that can be

achieved. This is an excellent technique for visualising leaf surfaces

in their native, hydrated form but necessitates the use of a SEM with

specialised capabilities. It is best suited for low-resolution images of

plant surfaces (up to 5000× depending on the samples). Despite its

inherent disadvantages, surface images taken by this technique may

serve as useful low-magnification controls to avoid misinterpretation

of the surface structure by artefacts introduced from dehydration

techniques. The technique also has the potential to be fruitfully used in

studying insect taxonomy, host: pathogen interactions and elemental

deposition in plant tissues.

Fig. 9. Chenopodium leaf sample processed by air-drying and observed under a FESEM. (a) Salt gland with waxes clearly visible on surface, compare this with Fig. 1B where waxes were completely obscured when ESEM was used; (b) base (stalk) of the gland left on leaf surface after the gland is physically detached; (c) gland surface wax micro-morphology; (d) leaf surface wax micro-morphology. Note the high resolution and brightness achieved at low electron dose (3 kV) using this technique.

(a) (b)

(c) (d)




LTSEM followed by cryofixation of samples may prove useful as high

magnification controls, but this technique is not entirely artefact-free.

In addition the technique necessitates the use of a specialised cryo-

chamber attached to the microscope. If a cryo-stage is not available,

samples processed by simple air-drying and examined using appropriate

beam and probe current conditions may provide high-resolution

wax images (up to 50,000× magnifications) as demonstrated in the

current study. CPD following chemical fixation with glutaraldehyde

and/or osmium tetroxide gave excellent preservation of internal

leaf structures, but surface preservation was less than ideal for

many samples. Simple air-drying following chemical fixation (with

glutaraldehyde and osmium tetroxide) is not recommended at all for

sample preservation because the fixatives caused severe distortion of

leaf tissue by negatively influencing the drying process. Fixation with

methanol before carrying out the drying process provided some good

preservation, but the technique needs to be refined further for specific

plant species.

There is no universal method for plant tissue processing for scanning

electron microscopy. Plants vary in their tissue characteristics and are

relatively difficult to preserve in their original form as compared to

animal or insect tissues. Specific techniques need to be developed and

tested for a specific objective. The technique that can be employed

for a plant tissue is dictated by the surface features to be preserved,

the resolution and magnification to be achieved and availability of

processing equipment and the capabilities of electron microscope

available—a case of horses for courses.

AcknowledgementsThe project was funded by New Zealand Foundation for Research Science

and Technology. Thanks to Lloyd Donaldson and Adya Singh for useful

technical advice on microscopy issues and operations; Jerzy Zabkiewicz

for suggestions on freeze-drying techniques. The assistance provided by

Catherine Hobbis and Bryony James for the use of FEI Quanta ESEM is also

gratefully acknowledged.

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Originally published in the Elsevier journal Micron. doi:10.1016/j.micron.2008.05.006


sample preparation of sem for plant smaples

Engineering

specific techniques

sample preparation methods

specific leaf sample

plant tissues

conventional microscopes

similar techniques

environmental sem microscopes

available electron microscope