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Why shouldn't I clean microscope slides with a paper towel?

Why shouldn't I clean microscope slides with a paper towel?


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I've recently bought a microscope and been looking at a lot of bacteria and such, and have quickly run out of clean slides. I've seen videos and articles saying how you shouldn't clean microscope slides with a paper towel and to use an ultrasonic cleaner instead. I don't have access to an ultrasonic cleaner, so I am wondering what is wrong with cleaning them with a paper towel?

Also if anyone knows of any good liquid solutions for cleaning slides that information would much be appreciated.


The main reason to avoid paper towels, tissues, and etc. is that they shed lint (paper fibers) which will be left behind on your slide and eventually interfere with your prep. They look small to the naked eye, but they are not small compared to the prep on your slide.

The other issue with paper towels and etc. is that there might be some particulate matter in the paper that's hard enough to scratch glass or optical coatings. This is more of an issue for optics - especially coated ones - than for cheap microscope slides, but it's something to consider.

If you're not mounting permanent preps then ultrasonic cleaning is probably a bit of overkill - although I have a surplus sonicator and it does a great job on a lot of tasks. It's generally sufficient to rinse the slide with distilled or deionized water and then follow up with (clean) alcohol. Optical purists will say spectroscopy grade methanol, but for home use any reasonably pure MeOH, EtOH or iPrOH will probably do. The main concern, again, is leaving behind particulates or residues. You can allow the slides to air dry if you're using pure alcohol, or use a piece of lens tissue (lint free paper tissue) to help them dry faster. I've also used Kimwipes in a pinch, but they're not truly lint free so not ideal. They're still much better than a tissue or paper towel.


In Biology, the compound light microscope is a useful tool for studying small specimens that are not visible to the naked eye. The microscope uses bright light to illuminate through the specimen and provides an inverted image at high magnification and resolution. There are two lenses that magnify the image of the specimen &ndash the objective lens on the nosepiece and the ocular lens (or eyepiece). To determine the total magnification of the specimen, you must multiply the objective lens magnification with the ocular lens magnification.

Scientists and technicians often use light microscopes to study cells. Prokaryotic cells are very simple and lack a nucleus or membrane bound organelles and are small in size. On the other hand, eukaryotic cells are more complicated in that they contain a nucleus and many specialized organelles. A cell&rsquos structure dictates its function thus, each eukaryotic cell looks very different from the next. This is why a cardiac cell looks completely different from a neuron (brain cell).

It is very important to learn how to handle and use a microscope properly. Review the following rules and tips for using and handling your microscope.

Figure 1. Labeled parts of a microscope.

General Rules

  • Always START and END with the low power lens when putting on OR taking away a slide.
  • Never turn the nose piece by the objective lens.
  • Do not get any portion of the microscope wet - especially the stage and objective lenses.
  • Use only lens paper to clean microscope lenses.

Cleaning the Microscope

If needed, obtain a small square of lens paper (and ONLY lens paper) and gently wipe the microscope lenses directly across, in this order:

  1. the lower surface of all the objective lenses
  2. the ocular lens
  3. the condenser lens and the light housing

Washing Brand New Slides

Place a small drop of cleaning solution on each microscope slide. This can be dish washing fluid, or it can be a more specialized cleaning solution for slides, such as an ethyl alcohol solution.

Apply the soap uniformly across both sides of the glass with something that won’t scratch the slide, such as a lint-free microfiber towel.

Rinse the slide thoroughly using warm running water. Continue until all of the cleaning fluid is gone, including any extra bubbles that appear.

Blot the slide with a paper towel until it is dry. Alternatively, you can dry the slides with microfiber towels. Make sure that the towel you use is clean for each new slide. You may have to switch to a new towel after a certain number of slides.

Place each finished slide back into the slide case. Each case will typically carry 25 slides. Make sure that each slide goes into its proper place. If you try overloading the case with more slides than it can take, the slides could bang against each other and crack.


Microscope Slides Preparation

Objects magnified under compound microscopes are mounted onto microscope slides. Made of glass or plastic, slides are approximately 1x3 inches and between 1mm-1.2 mm thick.

Multiple methods of preparation allow for advanced viewing of inorganic and organic objects.

Flat and Concave Styles

The most basic of all microscope slides is a flat rectangular piece of soda lime glass, borosilicate cover glass or plastic, with ground edges.

All corners are a sharp 90-degrees and, along with a rough outer edge, can cause minor finger cuts if not handled with care.

The top and/or bottom edges of a slide can be frosted, enabling easy marking for sample identification and/or orientation. The etched frosting keeps all pen marks safely away from the sample and a selection of frosted colors provides additional means of categorization.

Rounded safety corners to prevent accidental cuts as well as beveled edges with clipped corners ideal for blood samples are options available for both generic and frosted slides.

Concave microscope slides contain one or more surface depressions ideal for liquid solutions and larger specimens. These more expensive microscope slides can be used without a cover.

Some manufacturers produce plastic chambers with a predetermined number of slides with covers.

Filled calibrated wells or flasks are viewed quickly without preparing or clipping individual slides to the microscope stage, making this especially useful in sediment studies, such as urine analysis in addition, some tray designs can be placed in an incubator or refrigerator, allowing for the study of cultured samples.

Additional Features

Some cells and tissues cannot adhere to a plain glass surface and require a positive charge or surface modifications.

Saving time and money, electrostatic charged slides are a popular choice for researchers of histology, cytology and pathology.

Surfaces treated with biological reagents can make a slide water-proof, resistant to certain chemicals and reduce instances of cross-contamination.

Additional variations to microscope slides include:

Etched grid system or graticule

  • Enables researcher to monitor and communicate area(s) of interest
  • Aids in hand sketching
  • Helps geographical plotting
  • Estimates size and scale
  • Side-by-side comparisons, including sample to control
  • Reduces risk of cross-contamination
  • Rarely used substitute for glass
  • Less prone to dust, scratching
  • Prevents glare

Cover Slips

Almost always made of borosilicate or silicate glass, cover slips hold samples in place and protect them from inadvertent movement and contamination.

It also protects the microscope, preventing direct contact between the sample and lens as well as accidental leakage of water-based preparations.

The thin, transparent cover glass is usually square and available in types Number 1 and Number 2.

Suited for high-resolution microscopy and oil immersion preparations , Number 1 covers are .13-.17mm thick. Number 2 covers, .17-.25 mm, are designed for general purpose.

Less frequent characteristics include rectangular shapes, alternative materials such as quartz and certain types of plastic, etched lines or grids and additional thicknesses.

If not creating a permanent slide with glue and/or sealant, cover slips can be removed and sterilized for multiple reuses.

Preparation Techniques: Dry Mounts, Wet Mount, Squash, Staining

The main methods of placing samples onto microscope slides are wet mount, dry mount, smear, squash and staining.

The dry mount is the most basic technique: simply position a thinly sliced section on the center of the slide and place a cover slip over the sample.

Dry mounts are ideal for observing hair, feathers, airborne particles such as pollens and dust as well as dead matter such as insect and aphid legs or antennae. Opaque specimens require very fine slices for adequate illumination.

Since they are used for primarily inorganic and dead matter, dry mounts can theoretically last indefinitely.

Used for aquatic samples, living organisms and natural observations, wet mounts suspend specimens in fluids such as water, brine, glycerin and immersion oil. A wet mount requires a liquid, tweezers, pipette and paper towels.

  • Place a drop of fluid in the center of the slide
  • Position sample on liquid, using tweezers
  • At an angle, place one side of the cover slip against the slide making contact with outer edge of the liquid drop
  • Lower the cover slowly, avoiding air bubbles
  • Remove excess water with the paper towel

Although wet mounts can be used to prepare a significantly wide range of microscope slides, they provide a transitory window as the liquid will dehydrate and living specimens will die.

Organisms such as protozoa may only live 30 minutes under a wet mount slide applying petroleum jelly to the outer edges of the cover slip creates a seal that may extend the life of the slide up to a few days.

In addition, larger protozoan such as paramecium may be too large and/or move too quickly under the wet mount.

In these circumstances, adding ground pieces of cover glass to the water before the slip layer will create added space and chemicals or strands of cotton can be added to slow the movement of paramecium, amoeba and ciliates.

Smear slides require two or more flat, plain slides, cover slips, pipette and tissue paper:

  • Pipe a liquid sample such as blood or slime onto a slide
  • Using the edge of the second slide, slowly smear the sample creating a thin, even coating
  • Put a cover slip over the sample, careful not to trap air bubbles
  • Remove excess liquid

Ideally, smears should dry naturally in an environment of moderate, steady temperature.

The angle of the smearing slide determines the length of the smear a steeper angle creates a shorter smear. For samples such as blood, begin by backing the smearing slide into the sample and then push across the slide, pulling the blood in the opposite direction to create a smooth layer.

A thicker slide can be created with two drops, but only with the blood of mammals as the erythrocytes lack a nucleus allowing cells to be amassed in multiple layers.

Read further about Blood Smears - process and technique, possible presence of artifacts

Designed for soft samples, squash slides begin by preparing a wet mount place lens tissue over the cover glass gently press down, careful not to destroy the sample or break the cover glass, and squash the sample remove excess water.

A variety of methods exist for staining microscope slides, including non-vital or in vitro stains of non-living cells and vital or in vivo stains of living tissue. Staining provides contrast through color that reveals structural details undetected in other slide preparations.

Staining solutions such as iodine, methylene blue and crystal violet can be added to wet or dry mounts.

  • Add a drop of staining solution on the edge of one side of the cover slip
  • Position the edge of a paper towel on the opposite end
  • Allow dye to be pulled across the specimen

Stains are especially useful in the fields of histology, virology and pathology, allowing researchers to study and diagnose diseases, identify gram positive and negative bacteria as well as examine detailed attributes of a variety of cells.

Prepared Slides

Especially useful for educational purposes and for those who do not want to undertake the laborious process of creating slides, prepared microscope slides are available in all areas of science, including:


Why shouldn't I clean microscope slides with a paper towel? - Biology

Microscopes often represent a significant investment of funds and are sophisticated optical instruments that require periodic maintenance and cleaning to guarantee production of high-contrast images equal to the quality of the optical, electronic, and mechanical components. When neglected by exposure to dust, lint, pollen, and dirt, failure to remove immersion oil in a timely manner, or when expensive objectives are abused, optical performance can experience a serious decline that increases over time.

A microscope that remains unused for a lengthy period of time can accumulate dust and debris from the air (a condition that is only aggravated by leaving the instrument uncovered), which can lead to deterioration of image quality even though the instrument may be practically new. Proper use and regular maintenance of the microscope's mechanical components are equally important to prevent impairment of operation and eventual damage to the entire mechanical integrity of the instrument. The best instrument covers are designed to provide maximum protection from airborne contaminants for specific microscope types, as they are typically configured with their common attachments (see Figure 1). Even when carefully covered for protection during periods of inactivity, microscopes that are used regularly are subject to build up of contaminants. Some of these are unavoidably introduced from the environment and others by the microscopists themselves, especially in areas where the hands, eyelashes, and even moisture from breathing contact the instrument over time.

Blemishes such as dust, lint, and smudges on the optical components, as well as scratches, pinholes, and striae in the lenses, filters, prisms, mirrors, and faceplate of the image sensor, tend to degrade overall microscope performance. The objective front element illustrated in Figure 2 exhibits a variety of particulate contamination, as well as severe scratches that seriously degrade its performance. When the optical elements at or near the conjugate field (image) planes are dirty, damaged, or defective, artifacts are likely to appear in sharp focus superimposed on the specimen image. Ironically, the higher the quality of optical components, such as the condenser and collecting and relay lenses, the more these blemishes interfere and contribute to optical noise .

After a source of optical noise is localized to a given component (by turning or shifting the suspected components in turn), the dirt may be removed by a variety of procedures discussed in detail in subsequent paragraphs. The utilization of immersion oil is essential in maximizing microscope optical performance, but its improper use or the failure to immediately remove the oil after each use constitutes the most serious contaminant that must be dealt with in instrument maintenance. Because immersion oil is a known substance intentionally applied to the microscope to enhance optical performance, its clean-up is discussed separately from the removal of other debris that inadvertently accumulates on either the mechanical or optical microscope components.

Routine Removal of Loose Particulate Matter

If the microscope has been idle and uncovered for a lengthy period of time, a significant amount of debris accumulation has probably occurred. In the typical laboratory environment, a surprising amount of particulate material can be seen to accumulate on an objective or other component that is left uncovered on a bench for even a short period, such as overnight. Because such debris is often highly abrasive, it must be removed from the microscope frame and mechanical parts with care, using a small vacuum cleaner or by dabbing with a moist paper towel. Dirt that is non-adherent may be removed from less delicate lens surfaces by gentle brushing with a clean camelhair brush. Figure 3 illustrates two of the basic cleaning tools commonly used in microscopy. Alternatively, an air blower or compressed gas duster can be employed, but it must be assured that no oil or similar spray is released from the compressed gas can.

Several manufacturers produce oil-free compressed gas cylinders that are ideal for dusting glass surfaces if appropriate precautions are followed (see Figure 4). The common small portable cans of compressed gas must absolutely not be tipped or shaken while spraying in order to avoid release of cold liquid propellant. Although it is difficult to resist the almost reflexive tendency to blow away dust by mouth when it is noticed on lenses and other areas of the microscope, this should be avoided. One should never attempt to blow the dust off lens surfaces with a strong breath because doing so risks spraying the lens surface with droplets of saliva that can mix with dirt to produce an abrasive slurry. A deliberate and systematic cleaning protocol is recommended for thorough contamination removal, and appropriate techniques are detailed in the following sections. While it is often suggested that a regular maintenance schedule be followed at periodic intervals, the necessity for cleaning is dictated by the use of the instrument and by the effectiveness of preventive measures taken to avoid build up of debris. Delicate components should only be cleaned when necessary, as most scratches and other damage to optical surfaces result from improper attempts to clean them.

Proper Use and Removal of Immersion Oil

Following proper procedures in the use of immersion oil will significantly ease the task of removing the oil from microscope components before it causes damage. It is important to recognize that immersion oils are not inert with respect to either optical or mechanical microscope components, and if left in contact with the instrument, oil will penetrate into gears and sliding mechanisms and into crevices between lens elements and their mounting structures, with the potential to cause irreversible damage. Even when employed properly, immersion oil must be removed immediately after use to prevent its accumulation in unwanted areas of the microscope, as well as to avoid optical degradation from dried oil residue on the objective. Oil that has been stored for more than one or two years may not perform optically the same as fresh oil, and a potential increase in viscosity often makes it more difficult to remove. Consequently, containers of immersion oil should be labeled with the date received, and discarded when necessary.

The full utilization of the microscope optical system numerical aperture when immersion objectives are used requires a double oiling technique in which immersion oil is applied to both the top and lower surfaces of the specimen slide. Placing immersion oil in the gaps between the objective and slide and between the condenser and slide provides a homogeneous optical medium from the condenser, through the specimen (in an appropriate mounting medium), and into the objective. Although the viscosity of immersion oil minimizes any immediate migration into unintended locations, if it is not removed promptly and is allowed to accumulate, the effects of gravity and capillary forces will ultimately result in the oil moving into parts of the substage mechanism and microscope stand, and perhaps even into the objective. This accumulation may not be readily visible, and can go unnoticed until mechanical or optical problems become severe enough to require service by a microscope repair facility.

Correct utilization of immersion oil requires placing a single drop on the top lens surface of the substage condenser and another single drop on the top of the specimen slide. The condenser is then raised just to the point that the oil drop contacts the lower surface of the slide, and the objective front lens is brought into contact with the oil drop on top of the slide. It should be stressed that the oil immersion technique is only to be used with a condenser equipped with an immersion-type top lens, and with immersion objectives. Any attempt to improve the performance of a dry objective by application of immersion oil will likely result in its destruction, as such objectives are optimized optically for use in air, and are not sealed against the intrusion of fluids into the lens barrel.

After each specimen has been studied, the immersion oil should be completely removed, even if additional slides are going to be observed. While it seems expedient to simply add additional drops of oil when changing to the next specimen, this practice results in excess oil accumulating on the microscope, which will eventually find its way into damaging locations in the substage assembly and even the microscope stand. Only a single drop of oil at each specimen-optical interface can be accommodated without producing contamination that may be impossible to remove without complex disassembly or factory servicing of the instrument.

Immersion oil is most safely removed using only lens tissue, without employing any solvents. After moving the stage away from the objective, and lowering the condenser away from the slide, the slide can be removed from the stage and set aside for subsequent cleaning. With most microscopes the objective that requires cleaning is most easily accessed by swinging the lens turret to position the objective toward the front of the microscope. Lens cleaning paper that is specifically for use on high quality optics must be employed, and it should be stored in a covered container to prevent contamination with airborne particulates. A folded piece of lens tissue is drawn across the objective front lens to absorb the oil, and repeated with a new area of the tissue. This gentle wiping of the lens surface should be repeated, with as many tissues as required, until no oil streaks are seen on the tissue, and each tissue discarded immediately to avoid inadvertently reusing contaminated tissues on the objective. The folded tissues can be held under light tension with two hands while wiping, or pulled across the lens like a paper swab.

Direct pressure from the fingers should never be applied to the glass lens surface through the paper in order to minimize the possibility of scratching the lens if any particulates are present on the tissue. It should be emphasized that using a number of fresh lens tissues is essential to the success of this procedure, and the natural tendency to minimize "waste" is definitely misdirected economy considering the relative cost of lens tissue compared to the potential of damaging an expensive objective. If 20 tissues are required to clean an optical component, then that many should be used and discarded without hesitation.

When no residual traces of immersion oil are apparent on the final tissue paper, another tissue should be employed to wipe the lens with moisture from the breath. As cautioned previously, one must not blow through closed lips onto the lens, but should breath gently on it with the mouth open, so that no saliva droplets are expelled. If possible, the mouth should be positioned beneath the level of the objective to further reduce any possibility of droplets landing on the lens. With moisture condensed from the breath as a lubricant and solvent, a fresh piece of lens tissue is used to wipe the lens surface in a circular motion. An effective method of preparing lens paper for this cleaning is to fold all four corners of a piece of tissue together, leaving the untouched center of the tissue bulging out. The corners can be twisted together slightly to form a stem for handling the tissue. When the tissue is held by this stem, and wiping performed with the puffed-out tissue center, the force that can be applied to the objective is limited by the springiness of the tissue. Circular wiping motion can be applied in this manner, with very little direct force on the lens surface.

The procedure of breathing on and wiping the objective front lens should be repeated several times with a new tissue each time. With high-magnification objectives, having very small front lens elements, the lens paper can be twisted into a sharper point if necessary, taking care not to touch the portion of tissue applied to the lens. The spring effect of the paper can still be exploited to limit the force that can be applied to the lens surface when cleaning. Removal of immersion oil without removing the objective from the microscope assumes that the structure of the instrument does not restrict access to the objectives. In the latter case, the objective must be carefully removed from the nosepiece and placed on a suitable protected surface on the lab bench for cleaning. In any instance, objectives that are regularly used should be removed (one at a time) for a thorough cleaning at periodic intervals. This allows each to be more carefully inspected, as described in a following section, for signs of any type of accumulated contamination. Figure 5 demonstrates cleaning and inspection of an objective that has been removed from the microscope. The small front element of the objective can be effectively cleaned with a tissue formed into a point (Figure 5(a)), and the effectiveness of the cleaning evaluated under magnification using a loupe or inverted ocular (Figure 5(b)).

Ideally, the removal of immersion oil from the objective is successfully accomplished only through the mechanical application of lens tissue, and a similar procedure is then applied to the condenser top lens. It may be advantageous to remove the top lens of the condenser to facilitate cleaning, especially if removing it minimizes the likelihood of dispersing oil into other parts of the condenser body. The procedure described for cleaning the front of the objective should be repeated with the condenser lens that was oiled, and the body of the condenser inspected for any stray oil, which must be removed. Following cleaning of the optics, immersion oil should be cleaned from both surfaces of the specimen slide using laboratory tissues (brand names such as Kimwipes or Micro-Wipes ). It is not necessary to utilize lens tissue for removing oil from larger areas such as specimen slides, or from other portions of the microscope base or stand. All such areas on the instrument should be routinely checked for any traces of immersion oil, which if found, may be removed with laboratory towels or soft cotton cloth.

Inverted (tissue culture) microscopes present special problems with regard to the use of oil-immersion objectives because spilled or migrating oil can very easily intrude into the interior of the objective at the juncture between the body and the telescoping spring-mounted front lens barrel. If oil is allowed to accumulate, it can conceivably flow, under the force of gravity, even into the objective turret or nosepiece. Specially designed higher-viscosity immersion oils are available for use with inverted microscopes, and should be employed to prevent migration of oil from the objective front element.

Hazards of Solvent Cleaning

Numerous publications by respected authorities in microscopy, including several microscope manufacturers, recommend the use of various solvents as aids in removing immersion oil from objectives and other optics, as well as for routine removal of other contaminants. While this may simplify and accelerate the cleaning process, the variations in lens construction and the materials used in other microscope components, as well as the health and safety hazards presented in using most of the applicable solvents, make it inadvisable to recommend their general use. Extreme care must be exercised in applying solvents to components that may be irreparably damaged if solvent migrates into internal areas or if it is applied in excess and remains in contact with the surface for too long before evaporating. Many cleaning procedures that have been used successfully for decades have become unacceptable today for a variety of reasons, including additional knowledge of health and safety hazards associated with the solvents for organic non-polar compounds used in immersion oils. The issue of the use of solvents is complicated, and is confused by contradictory recommendations in the scientific literature, as well as by differences in manufacturers' technical publications. Some of the considerations relevant to solvent cleaning are discussed in more detail in the following sections.

In the past, solvents have been routinely employed for nearly any cleaning task in microscopy, and particularly for removal of immersion oil. Potential problems associated with solvent cleaning are sufficiently serious that the best current approach in cleaning the microscope is to use solvents only when absolutely necessary, essentially as a last resort rather than a first step. Information provided in instruction manuals of microscope manufacturers exemplifies the difficulty in selecting a cleaning solvent when one is required. Some manufacturers have for years warned specifically against the use of alcohol as a lens-cleaning solvent, while others recommended ethanol and mixtures of ethanol with other solvents. An ideal solvent would be miscible with organic non-polar compounds, not highly flammable, sufficiently volatile to evaporate quickly leaving no residue, and be non-hygroscopic and non-toxic. Most solvents that have been routinely used historically fail one or more of these criteria. With optics allowing use of alcohols, a mixture of ether and ethanol (50:50 by volume) is effective, as is the modified mixture of ether, ethanol, and chloroform (48:48:4 by volume), but both are dangerously flammable or explosive, and produce toxic vapors.

One of the most significant dangers with many of the solvents proven effective for cleaning microscope optics is that they have the potential to dissolve the cements utilized in lens assembly (as do the immersion oils themselves if allowed to remain on the optics). In the past, benzene was regarded as a highly effective lens cleaning solvent, but always required great caution to limit contact with the lens for no more than a second or two, due to the high solubility in benzene of balsam and some other cements used for lens mounting (and for mounting coverslips on specimen slides). The high volatility of benzene is an advantage in this regard, but the material is also highly flammable and toxic. It is now known that benzene is readily absorbed through the skin, and this as well as inhalation of the vapors can cause liver damage. As a consequence of the numerous hazards, benzene should never be used for cleaning. Xylene has been widely utilized for years, and is considered a less aggressive solvent than benzene, but because of its lower evaporation rate, residual liquid may be more likely to penetrate and damage a lens unless the xylene is used very sparingly. Xylene is, however, highly flammable, toxic and carcinogenic, and may cause skin contact sensitivity. Although alcohol and xylene are widely recommended as lens cleaning solvents, they are also named as being harmful to both the mechanical and optical components of many microscopes. The finish on portions of the microscope stand and the materials used in a number of the parts themselves can be severely damaged by exposure to either material.

Because of the variation in solvent recommendations, and the likelihood that some of the materials used in the instrument components are not known to the user, it is prudent to restrict use of any solvent to an absolute minimum. Optical components should not be immersed in any solvent, and cleaning tissues should only be moistened, never saturated, with a cleaning solution. Minute gaps commonly exist at the glass-metal junctures of an objective front element, allowing the possibility of solvent migration into the interior of the optical component if excessive solvent is applied. Depending upon its composition, the optical cement used to join lens element combinations in objectives is commonly soluble in one or more of the solvents, alcohol, xylene, and acetone. The result of solvent penetration between lens elements is illustrated in Figure 6, in which the partial separation of cemented lens groups has occurred. Although most modern optical cements are not readily affected by xylene, some older objectives utilize cements that are totally soluble in xylene.

Alternative Cleaning Materials

Several alternatives to hazardous solvents have been found to be effective in microscope cleaning, and a variety of cleaning agents, as well as cleaning materials, are recommended by different microscopists and manufacturers (see Figure 7 for examples). Safer alternatives to xylene have been widely pursued, in part because that solvent is commonly used in histopathology and cytology laboratories as a depariffinizing and clearing agent. The proprietary solvents Histolene and Histoclear are gaining popularity as replacements for xylene in microscopy laboratories, and have been found to be effective for instrument cleaning as well. These solvents are based on the naturally-occurring compound d-limonene , which is the major constituent of citrus peel oils and other ethereal oils, and which has been used extensively in the food and cosmetics industries for years. Although the limonene-based solvents require adequate ventilation and skin protection, they currently are thought to be safer overall than xylene. Pure distilled water is the safest cleaning fluid for any contamination that is water soluble if that is inadequate, commercial photographic lens cleaning liquids are very effective and are safe for precision optics when used sparingly. This type of cleaning agent consists primarily of water to which is added a small percentage of surfactant and alcohol. Commercial window cleaning products (such as Windex and Sparkle) are used by some microscopists, with no reported damage to optical components, and isopropyl alcohol is employed successfully by others.

Great care must be taken in choosing materials for applying water or other cleaning liquids to precision optical components. Although many products are marketed as being suitable for lens cleaning, and other materials give the subjective impression that they would not be harmful, the suitability of specific materials for delicate optics is not always obvious. As an example, the laboratory tissues marketed under the name Kimwipes have been shown to be suitable for lens cleaning, although they feel quite coarse to the touch. In contrast, typical facial tissues are processed to feel soft to the skin, but contain hard particulates that are definitely harmful to optical surfaces. Lens tissues are available in varieties that feel relatively stiff and hard-surfaced, with tight fibrous texture, and others that are loosely textured and very flexible. The softer type is generally preferable for delicate optics, even though these tissues tend to leave residual loose fibers following cleaning, which must be blown off with air. Freshly laundered pure cotton or linen fabric is recommended by some microscopists for lens cleaning, but with any material that is reused, it is essential that no detergent residues or particulates remain after washing. Not only is this not a trivial requirement to meet, it is also important to ensure that if manufactured cloths such as handkerchiefs are used, they are not hemmed or otherwise sewn with polyester or other abrasive thread.

A common recommendation in the past for performing lens cleaning was to wrap small portions of cotton wool around the tip of an orangewood stick (an oil-free wood) for use as a cleaning swab. This is no longer advisable, due to the fact that cotton wool such as that now sold by pharmacies in rolls typically contains some proportion of synthetic fibers, and is not as suitable for delicate surfaces as is 100-percent cotton wool. Cotton swabs that are untreated are still considered to be suitable, although these are wound into very tight buds at the factory, and before use it is wise to loosen some of the cotton at the tip of the swab with clean forceps (not the fingers, which will deposit skin oils) so that less force is applied to the surface being cleaned. Applicators made by attaching small pieces of clean chamois to orangewood sticks are commonly used by optical technicians, and these are commercially available or can be made-up in special sizes, as desired.

Basic Cleaning of Mechanical Components

The primary concern in maintenance of the mechanical components of the microscope are areas of the instrument which are unavoidably exposed to skin oils from the hands and moisture from breathing, and the stage area, which is subjected to a variety of contaminants during imaging sessions. In addition to the stage, other components to be cleaned include controls such as knobs, levers, and movable control rods, the body tube, and the stand. Because many of the microscope controls, such as focusing knobs, are ribbed or milled in a fine crosshatch pattern, skin oils tend to collect in these areas and attract dust, which can become tightly bound to the control. Cleaning may be required frequently on microscopes that are heavily used. An effective cleaning liquid may be prepared by adding approximately 10 percent alcohol, by volume, to a commercial glass and surface cleaning product. A piece of terry cloth toweling moistened with the cleaner should be used to remove contamination from the ridges of every control by wiping in the direction of the ridges, or in multiple directions on milled surfaces. Prepackaged moistened wipes for optical components provide an alternative method of applying a controlled amount of cleaning fluid, which may be effective for cleaning many microscope surfaces (see Figure 8). Each cleaned control surface should be dried with a clean piece of toweling.

As a microscopist works with the eyes adjacent to the oculars, the close proximity of the facial areas around the eyes and nose to the cooler surfaces of the body tube result in vaporized moisture and skin oils condensing on these microscope surfaces, leading to a significant amount of contamination. Additionally, the breath impinges on both the body tube and objective nosepiece, contributing further to the collection of airborne contaminants on the moist surfaces that result. The use of an air deflection shield, commonly referred to as a breathshield , on the microscope is effective in reducing this source of contamination by diverting the breath away from the nosepiece and microscope stand. The body tube and other parts of the instrument stand can be cleaned with soft cotton cloth lightly moistened with the surface cleaner referred to previously. It is especially important to clean the area around the eyepiece interocular distance adjustment mechanism, which is particularly prone to the build up of contamination. In order to avoid getting any moisture inside the eyepiece tubes, they should not be wiped with the moistened cloth near the top at the mating surface where the ocular rests. After cleaning the body tube, it should be dried with another piece of cotton cloth, and this dry cloth can be used to clean the top portion of the eyepiece tube and the outer rims of the oculars, taking care to avoid touching the glass lens surfaces.

The microscope stage is cleaned in a similar manner to the body tube, first with a moistened cloth, then with a dry one. Because of the variety of contaminants that may be deposited on the stage from specimens and from constant handling and manipulation, it should be cleaned after every use of the microscope. Care must be exercised in cleaning around the edge of the center opening in the stage, and contact should not be made with the underside of the stage where there may be exposed grease from bearing surfaces. Any cloth contaminated with the special grease used on the instrument stage should be discarded to avoid transferring it to other parts of the microscope, as it may be virtually impossible to remove.

The remainder of the microscope stand should be cleaned carefully with the same procedure of a moistened cotton cloth followed by a dry cloth, taking care to avoid optical surfaces or any area that might be subject to moisture penetration that could damage internal mechanisms or electronic circuitry. Following complete cleaning of the mechanical components as described, and carefully wiping up any liquid spills in the vicinity of the instrument, a small vacuum cleaner (see Figure 3), with a flexible hose and soft brush attachment, can be employed to vacuum up any loose material on the stand and table area around it. Extreme care should be taken to avoid touching any optical surfaces with the vacuum brush.

Basic Cleaning of Optical Components

A systematic protocol for inspection and cleaning of microscope optical components is essential for several reasons. Not only are the optics the most crucial components in image formation and recording, they are the most expensive, as well as the most delicate and most subject to damage. Inspection of optical surfaces with magnification, provided by a loupe or an inverted ocular, is an important first step in cleaning. Evaluating whether contamination is present and determining the type of material is important both because unnecessary cleaning is counterproductive and because certain types of contamination are not obvious without careful inspection. In particular, the front elements of the objective and condenser should be regularly inspected with a magnifier under reflected light by carefully positioning a light source at an angle to the surface being examined so that any debris can be seen. In troubleshooting a blurry or low contrast microscope image, it can be assumed that the most likely cause is a dirty front objective element, debris on glass surfaces near the imaging sensor, or a dirty coverslip. High-magnification objectives, with very short working distances, are especially vulnerable to contamination, and require frequent inspection.

The presence of even minor dirt or smudging on an objective, no matter what the nature of the material, produces the same effect, which is a reduction of image sharpness. This is true for particulate material and for contamination with perfectly transparent material such as immersion oil. Oil traces, including greasy fingerprints on a dry objective front element, interfere with the transmission of light rays through the objective in the same manner as would a damaged lens or one having an optical manufacturing defect. Inspection of the front objective element is the best way to determine whether contamination is present, and if so, what course of action is required for its removal.

The cleaning procedures described below apply only to exposed surfaces of the various optical components of the microscope. No attempt should ever be made to clean internal optical surfaces of most microscope components, and cautions are given in each of the following sections pertaining to specific components in order to emphasize potential damage that can be caused by not strictly following this advice. The basic cleaning protocol for optical surfaces, which is generally followed for all optical components, should be undertaken in steps, as follows:

Inspection of the Lens Surface - The optical component to be evaluated is removed from the microscope and placed on a laboratory towel or similar protective surface on the instrument table. Before any cleaning is attempted, the optical surface should be inspected with magnification under reflected light to determine the condition of the component. Particular attention should be given to the presence of any particulate material, which must be assumed to be abrasive, and removed before any other cleaning is done. Additionally, the presence of any films, smudges, or stains should be noted. A magnifying lens of 2-3x is appropriate for examining larger optics such as oculars and condensers, while the smaller lens elements of objectives require approximately 5x to 10x magnification for proper inspection. It is crucial that particulate matter be removed from a lens surface as the first step in cleaning, because any particle can be abrasive and result in scratches if it is moved across the surface with even the most gentle lens tissue.

Removal of Non-attached Particles - If any dust, fibers, or other particles are observed on the lens surface, an attempt should be made to remove them in the least aggressive manner possible, which is by gently blowing air across (not perpendicular to) the lens surface. The safest method of air dusting is to use a rubber bulb or balloon, such as the ones intended for use as ear and enema syringes for infants (an ear syringe is illustrated in Figure 7). The larger enema syringe is appropriate for larger optics such as eyepieces, condensers, and prisms the smaller ear syringe is better for small objective lens surfaces. The reason for not blowing directly toward the particle and surface is that this can force abrasive particles into delicate lens coatings, and possibly make them more difficult to remove, ultimately damaging the surface in the process. The cumulative effect of repeated abrasions of this type, though minor, can degrade the performance of the optic. Care is required to avoid touching the tip of the syringe to the lens surface. The best advice is to avoid any use of compressed air cans for lens cleaning. It is difficult with these to control the pressure of air impinging on the surface being cleaned, and there is always the risk of either extremely cold air or freezing liquid being expelled onto a lens surface and causing irreparable damage. Neither lens coatings, nor optical cement between lens elements can withstand localized freezing without damage. If, for any reason, a canned-air duster must be used, it should be stabilized in an upright position to avoid tilting (which will expel cold liquid), and fitted with a length of flexible plastic tubing to allow air to be directed in the desired direction onto the optical surface. It is far preferable to utilize the manual air bulb type duster to completely eliminate this risk.

Reinspection of the Lens Surface - Inspection of the lens again with a magnifier will reveal whether all particles have been removed, and if so, any contaminating films should be noted for subsequent removal. If particles remain after the initial air dusting, another attempt should be made to remove them with air alone. Any that are still present on another examination are most likely attached through direct interfacial tension between the particle and surface, or due to an intervening film of some type, and these must be removed before further cleaning of film contaminants can proceed.

Removal of Attached Particles - Particulate material that resists removal by air dusting alone is most likely held by a surface film through a minute contact area, and can be dislodged by delicately applying slight lateral force to the side of the particle. This procedure requires practice, and definitely must be done with adequate magnification to ensure that no damage is done to the lens. To devise a tool for nudging attached particles from their positions, a thin bamboo skewer or wooden toothpick can be cut off to a very fine square point with a razor blade. After breathing very gently onto the lens (with mouth open wide) to produce condensed moisture, which should loosen the particle from its adherent film, the point of the wooden tool is brought into contact with the side of the particle. It is gently nudged sideways, taking care not to touch the lens surface with the tool. This process is repeated for any other attached particles, and the small ear syringe is employed to blow across the lens surface to remove the freed material.

Reinspection of the Lens Surface - The lens is inspected again under magnification to determine if all particles have been removed. If any remain, the removal procedure is repeated, and the lens inspected again. When all particulates have been removed, if no additional contamination is present, the component can be reinstalled on the microscope. If any other film, streaking, fingerprints, droplets, or other contaminants are present on the optical surface, the following steps are performed.

Removal of Water-Soluble Films - Water-soluble materials can be removed from a lens surface using lens tissue and moisture produced by slowly breathing onto the lens. The tissue should be utilized in the manner previously described to limit the force applied to the lens, and never just rubbed on the surface directly with finger pressure. In addition to the puffed-out tissue technique, several other methods are suitable for limiting the force applied by the lens tissue. One that is effective for relatively small lens surfaces is to roll a folded lens tissue into a tight tube, and then to tear it in half forming two shorter tubes each having a frayed end. The frayed end of each tube is used to clean the lens surface. The tearing action not only should dislodge any particles on the paper in that area, but the torn end minimizes the force that can be applied to the lens. After gently and slowly breathing on the lens with the mouth opened wide to provide moisture, the lens is cleaned with a frayed tissue tube in a circular motion starting at the lens center and working outward toward the periphery. The tissue is discarded and the process repeated with additional torn pieces, until the lens appears clean or no more improvement is noted. The lens may not become completely clean if any contaminants are present that are not water-soluble.

Inspection of Lens Surface - The lens is again inspected using magnification, and if it is completely clean, the component can be returned to service on the instrument. If any film-like deposits or smudges remain on the lens, it is most likely a non-water soluble material, which must be removed with the following additional cleaning step.

Removal of Non-Water Soluble Films - Contaminants on an optical surface that are not readily removed with water (other than immersion oil, discussed previously) require an additional cleaning component. One of the safest materials that is effective on deposits of this type is one of the commercial lens cleaning fluids for precision optics, which are usually composed of distilled water to which small proportions of a surfactant and alcohol are added (see examples in Figure 7). A very limited amount of fluid should be utilized, and it should never be applied directly to the lens surface. An effective means of controlling the amount of fluid allowed to contact the lens is to use a cotton swab to which is applied a very small drop of cleaning solution. The tip of the cotton bud should be inspected for any particulates that are present, and as described previously, the tightly wound cotton can be loosened slightly by pulling the tip with clean tweezers or teasing out some of the cotton with a needle. The lens can be cleaned by lightly applying the swab in a circular motion starting at the lens center and moving out. As an alternative to the cotton swab, a soft lens tissue may be twisted into a point, being careful to not touch it at the end, and used as discussed regarding removal of immersion oil. When employing a cotton swab in this manner, extreme care must be taken to limit the force applied to the lens surface, and this technique should never be employed except as a final cleaning step immediately after complete removal of particulate materials.

Still another effective method of utilizing a lens cleaning fluid so that very little force is applied to a small lens, such as an objective front element, is illustrated in Figure 9. With the component resting on a soft surface on the table, a single drop of cleaning fluid is placed on a folded tissue, and while supporting the tissue with both hands, the drop is brought into contact with the lens surface. The tissue is then drawn horizontally over the lens surface, which will leave a streak of fluid on the tissue. There should be no attempt to force the tissue into contact with the lens in fact the surface tension between the lens and drop of fluid may make it possible to slightly pull the tissue away from the lens while moving it across, if this is not done with so much force that the tissue loses contact with the lens surface. The process should be repeated several times with a fresh drop of fluid and a new tissue each time. After cleaning with the moistened swab or tissue, the lens surface should be dried by repeated application of several torn lens tissue tubes, discarding each after use. An indication of the success of the cleaning can be obtained by breathing slowly on the lens to moisten it, noting whether the moisture film is even and without disruption. As a final step, the moisture is removed by wiping in a circular motion with a lens tissue tube.

Final Evaluation of Lens Surface - Inspecting the lens surface with magnification is the final step in determining whether the component is completely clean before replacing it on the microscope.

Notes and Cautions on Cleaning Specific Components

Modern, highly-corrected objectives may contain over 15 individual lens elements, some joined by optical cement into compound lens groups, which are assembled at precise separation distances within the objective barrel. Objectives should never be disassembled in an attempt to clean internal lens surfaces, or for any other reason. The component lens elements are precisely centered optically, and assembled with a precision that cannot be duplicated outside of the manufacturer's factory setting, and any attempt at disassembly will undoubtedly result in a damaged objective. Even if access to internal surfaces were possible, they could not be successfully cleaned without damage, due to the fragility of the anti-reflection and other lens coatings that are commonly utilized. Most precision lens surfaces employ one or more interference-film coatings that may be only a few atomic layers thick. These coatings are protected by hardened protective layers on external lens surfaces, to enable them to tolerate normal cleaning procedures, but the coatings on internal surfaces are much softer and very easily damaged.

Under no circumstances should the rear objective lens element be cleaned, other than to blow off dust that settles there with the ear-syringe blower. Due to the construction of the objective, which makes access to the rear element difficult, attempts to clean the rear element risk introducing tissue fibers or other contamination into the interior of the assembly. The interior can be checked for contamination by looking through the objective from the front, with a light source (such as a bare lamp) positioned close to the rear element. Unfortunately, if internal contamination is present, it can only be removed by qualified service centers.

One previous caution that bears repeating is the importance of not applying excessive pressure to the front lens surface of an objective. The front element of higher-magnification objectives is a very small, partially hemispherical lens that is held in place by minimal contact with the lens assembly. The large amount of metal surrounding the small glass element gives these objectives a robust appearance that is deceptive, as the front element can be easily moved out of alignment by excessive pressure, resulting in a damaged objective. Furthermore, even if the element is not forced out of alignment, applying too much pressure during cleaning or through accidental contact can produce minute gaps at the juncture between the lens and the surrounding metal barrel, causing oil or cleaning fluids to be drawn by capillary force into the objective interior, destroying the objective.

The top lens of most condensers is removable, and cleaning involves application of the basic optical cleaning procedure to the top and bottom surfaces of the top lens, as well as to the top lens surface of the middle assembly. The component parts of the condenser body should absolutely not be disassembled. They are assembled with similar precision to objectives, and cannot be realigned outside of the manufacturers' facilities. The same is true for phase contrast elements and differential interference contrast prisms, as well as for polarizing devices that are components of some condensers. These elements must be realigned at the factory if disturbed, and should never be removed or disassembled. Because of its location, the substage condenser collects a variety of contaminants, and must be cleaned more frequently than other components. Due to its relative inaccessibility on most microscopes, the condenser usually requires removal for proper cleaning, and should be handled with the same care given to objectives.

After removal of the condenser from the microscope, the top element can usually be removed by unscrewing. If a filter carrier is present beneath the condenser, it should be swung away from the condenser and any filters removed for subsequent cleaning. The surface of the top element may be contaminated with both particulate and film deposits, and is cleaned following the basic protocol of first removing particles, and then films or smudges. The lower surface of this lens will typically only have particulate debris, but should be inspected with a magnifier to confirm this, and cleaned accordingly. The next step is inspection and cleaning of the upper surface of the middle optical section of the condenser that is exposed when the top element is removed.

The condenser should next be turned over so that the bottom surface of the lower lens assembly can be inspected and cleaned if necessary. The primary caution at this stage is to avoid damaging the iris diaphragm if one is utilized beneath the lower optical combination of the condenser. If present, the diaphragm must be opened completely to allow access to the bottom lens surface and to protect the blades of the diaphragm. These blades are extremely fragile and should not be cleaned, touched, or exposed to any liquid. If opening the diaphragm does not retract the blades completely into the rim of the assembly, do not attempt to clean the lower lens by reaching through the iris opening, as damage to the diaphragm is likely. Cleaning of any filter removed previously must be done with the same care as exercised with other optical components. Interference filters are constructed utilizing very thin vacuum-deposited films similar to anti-reflection lens coatings, and filters of this type are commonly utilized in the condenser assembly and elsewhere in the microscope optical path. Particular caution must be used in handling and cleaning of such filters to prevent damage to the thin coatings.

The optical trains of modern microscopes contain a number of precision prisms and front-surface mirrors, most of which are housed within the microscope base and stand. As a general rule, none of these components should be cleaned unless they are accessible without disassembly of any part of the instrument. When internal components of this type are dirty, they require factory service to be cleaned without damage. The only exceptions are three external prism surfaces that are accessible in the body tube, and a mirror that is exposed (without any disassembly) in the base of some microscopes. Front-surface mirrors employ an unprotected reflective coating (usually silver) on the front of a glass base, and are very easily damaged. Removal of dust and fibers can be accomplished with gentle air dusting, followed by very gentle cleaning with lens fluid only when absolutely necessary. Cleaning with tissue should be done employing every effort to limit friction on the reflective mirror surface, which is easily abraded.

Binocular body tubes contain prisms for the right-eye and left-eye light paths that are precisely aligned using special collimating equipment, and no disassembly for cleaning should be attempted except by factory service facilities. Because the initial assembly is done under clean-room conditions to ensure a minimum of particulate contamination, any internal cleaning efforts would probably only introduce additional debris. The external prism surfaces that are visible when the eyepieces are removed from the body tube can be carefully cleaned by blowing off particulates, followed by use of cotton swabs that are softened on their tips as described previously for objective cleaning. Dust should be blown off with a large infant syringe after inverting the body tube so that dust falls away from the prism surfaces and out of the body tube. If further cleaning to remove smudges or films is required, it may be necessary to provide moisture for this procedure by breathing on the cotton swab tip instead of the prism itself, because of the recessed location of the prisms within the body tube. When the body tube is turned upside down, the lower opening reveals the third prism surface or an optical flat covering it, and this surface should be examined for signs of contamination and cleaned carefully if necessary.

Eyepieces require fairly frequent cleaning of their external optical surfaces, but do not generally become contaminated internally. The eye lens top surface is vulnerable to many types of contamination due to its proximity to the microscopist and its likelihood of collecting airborne particulates. Because the eyepieces are frequently removed for various reasons during use of the instrument, the lower field lens surface can become soiled and should also be examined for debris. Both of these lens surfaces should be cleaned as required following the basic protocol for optical components. In the rare circumstance that dust or fibers are seen in the interior of an eyepiece when it is inspected following external lens cleaning, it is possible in some cases to remove the eye lens in its mount and the field lens in it mount, and to clean the tube interior and the inner surfaces of the two lens components. They must be very carefully reassembled in their exact original configurations or the eyepiece will not perform properly. Particular care must be exercised with the finely threaded lens cells to avoid cross-threading the components upon reassembly. Note however, that under no circumstances should a Filar micrometer eyepiece, a measuring eyepiece containing an internal reticle, or any type of digital-readout eyepiece be opened or disassembled for any reason. Doing so will destroy the calibration, and require factory restoration.

Microscopes that are equipped with digital cameras may develop a degradation in captured image quality or exhibit image artifacts caused by accumulation of contamination either on filter elements that are sometimes utilized in the camera adapter or on the optical glass window that may be incorporated to seal the camera housing and protect the CCD or CMOS image sensor. In practice, if dark specks or similar in-focus artifacts are observed in digital images, and they are not in the specimen plane, their most likely cause is particulate contamination on the image sensor or an associated filter surface. Some digital cameras incorporate removable infrared filters in the camera system, while in others the required filtration is an integral part of the sensor window. Because of the variety of configurations encountered in scientific digital cameras, the manufacturer's recommendations regarding cleaning should always be followed. Some cameras, particularly those in which the sensor is cooled, are hermetically sealed, and the sensor is not directly accessible.

In general, the optical glass surfaces on sealed cameras should be inspected and cleaned, if necessary, following the standard cleaning methods for lens surfaces, always removing particulate debris before gently cleaning the glass surface with moisture from the breath, followed by lens tissue moistened with lens cleaning fluid for non-water soluble contamination. If the window is difficult to access with lens tissue (such as with torn tissue tubes), cotton swabs can be used provided that care is taken to limit pressure on the window surface. In some cameras, the sensor surface is directly exposed within the camera body, and is highly likely to attract dust and other debris. Special techniques may be required in cleaning the sensor to avoid static charge damage to the device, and the manufacturer's service personnel should be consulted for guidance on proper procedures.

Fungal Growth on Optical Surfaces

An especially serious problem that may plague microscope optical components is the development of fungal damage. Formation and growth of fungal colonies may occur rapidly in some climates, and when established on glass surfaces, it is unlikely that they can be removed before damage has been done to the surface. Unfortunately, fungal growth commonly occurs in the interior of optical components, and may be quite advanced before it is even noticed. At least one microscope manufacturer states that over 50 percent of deterioration in optical performance is attributable to cloudiness caused by certain fungus types. Although there are over 100,000 fungus species, two members of the genus Aspergillus are believed responsible for most lens deterioration. Optimum growth conditions for these fungi are relatively high temperature and high humidity, but they also are more adaptable to lower humidity levels than most other fungi. Figure 10 illustrates both the optimum and tolerable growth conditions for these fungi growing on lens surfaces, in comparison to the most favorable conditions for other common fungus species. Unfortunately, the conditions most conducive to proliferation of the lens-damaging fungi match very closely the most suitable environment for humans. This greatly complicates attempts to eliminate or inhibit growth of the fungi on optical components.

Fungi growing on lens surfaces reduce lens performance due to the lowered transmittance caused by the cloudiness, as well as by light dispersion from the thread-like filaments ( hyphae ) of the fungal colonies. Fungi growing on glass surfaces are not attached by roots and can be wiped off, but unfortunately, residual corrosion marks remain and the original lens performance cannot be recovered. The corrosion is a form of surface etching occurring when an organic acid produced by the fungus mixes with water vapor from the air that accumulates on fungus hyphae. Lenses with significant fungal growth usually must be replaced, since the only effective means to avoid fungal damage to optical components is to prevent its growth in the first place.

Favorable conditions for limiting the occurrence of fungal growth on surfaces such as lenses include a low-humidity environment, sufficiently low temperatures, good ventilation, and occasional exposure of the surface to sunlight. Climatic factors cannot be controlled completely, and the use of air-conditioning systems and dehumidifiers in warm and humid climates is beneficial and necessary, but does not eliminate the growth of the highly resilient fungus types, which can adapt to a wide range of conditions (see Figure 10). The strategy of storing optical components under desiccated conditions is sometimes suggested, but this is not an advisable practice, because extremely low (0 percent humidity) moisture levels can accelerate the breakdown of cements used to join optical elements. The geographical area in which the microscope is located determines, to a large extent, the seriousness of fungal growth as a factor in instrument care.

Figure 11 presents, in chart form, the seasonal variability of fungus growth conditions for a number of worldwide cities. It can be inferred from the chart that fungal growth is least likely in regions having consistently low humidity, or in regions that have relatively low average temperatures during periods of high humidity. In some climates, it is virtually impossible to inhibit fungal growth unless the microscope can be placed in a sterile environment. The major microscope manufacturers produce special versions of some equipment for use in tropical or other fungus-prone environments. Among preventive measures that have been developed are to enclose an antifungal chemical substance inside objectives, eyepieces, and the microscope base, and to improve the effectiveness of seals on any moving parts to minimize the entry of dust and fungal spores from the environment. The chemical is a solid substance designed to slowly sublimate and produce an antifungal vapor that is harmless to the microscope optical and mechanical components. The antifungal activity can be maintained over long periods of time by encasing the chemical in a material with only slight air-permeability, thereby strictly controlling the sublimation rate.

Benefits of Preventive Maintenance

The ideal microscopy room would be designed specifically for that purpose, and incorporate every mechanism available for limiting contamination by dust, chemical vapors, and other airborne contaminants, as well as isolating the instrument from acoustic and mechanical vibration and temperature variations. This ideal situation is seldom realized, and most microscopes are located in areas subject to a considerable number of environmental deficiencies. Some contamination is unavoidable, due to the rigors of daily use, but at the very least, the microscope should be protected as well as possible during periods of non-use by covering the entire instrument with a suitable cover. Instrument manufacturers and aftermarket suppliers offer a variety of specially designed dust covers (see examples in Figure 1). Of several types of plastic cover, those made of softer more flexible material are probably less prone to attraction of dust. Lint-free fabric covers are also available, and provide an effective dust barrier that can minimize the need for cleaning.

While the cost of a modern research grade microscope can range from approximately a few tens of thousands to several hundred thousand dollars, if properly used and maintained, the basic optical and mechanical components of the instrument can easily outlive several generations of microscopists. Only if the instrument is used correctly and maintained regularly is it capable of producing the best image data possible. Careless, incorrect operation and maintenance techniques not only result in unreliable and poor quality images, but cause productivity at the microscope to suffer, and the instrument's useful lifetime to be greatly reduced.

Thomas J. Fellers and Michael W. Davidson - National High Magnetic Field Laboratory, 1800 East Paul Dirac Dr., The Florida State University, Tallahassee, Florida, 32310.


Slide Mount Instructions

Before you start building your slides, make sure you have everything you will need, including slides, cover slips, droppers or pipets and any chemicals or stains you plan to use.

You will be using two main types of slides, 1) the common flat glass slide, and 2) the depression or well slides. Well slides have a small well, or indentation, in the center to hold a drop of water or liquid substance. They are more expensive and usually used without a cover slip.

Standard slides can be either plastic or glass and are 1 x 3 inches (25 x 75 mm) in size and 1 to 1.2 mm thick.

Wet slides will use a cover slip or cover glass, a very thin square piece of glass (or plastic) that is placed over the sample drop. Without the cover in place, surface tension would cause the droplet to bunch up in a dome. The cover breaks this tension, flattening the sample and allowing very close inspection with minimal focusing. The cover also serves to protect the objective lens from interfering with the sample drop.

MOUNTS

There are four common ways to mount a microscope slide as described below:

Dry Mount

In a dry mount, the specimen is placed directly on the slide. A cover slip may be used to keep the specimen in place and to help protect the objective lens. Dry mounts are suitable for specimens such as samples of pollen, hair, feathers or plant materials.

Wet Mount

In a wet mount, a drop of water is used to suspend the specimen between the slide and cover slip. Place a sample on the slide. Using a pipette, place a drop of water on the specimen. Then place on edge of the cover slip over the sample and carefully lower the cover slip into place using a toothpick or equivalent. This method will help prevent air bubbles from being trapped under the cover slip.

Your objective is to have sufficient water to fill the space between cover slip and slide. If there is too much water, the cover slip will slide around. Take a piece of paper towel and hold it close to one edge of the cover slip. This will draw out some water. If too dry, add a drop of water beside the cover slip. Practice this until you get used to it.

Wet mounts are suitable for studying water-bound organisms such as paramecium or bodily fluids such as saliva, blood and urine.

Section Mount

In a section mount, an extremely thin cross-section of a specimen is used. Using a microtome, cut a thin slice of your selected specimen such as an onion, and carefully set it on your slide. Then follow the instructions for a dry or wet mount. A stain can often be applied directly to the specimen before covering with a cover slip.

Section mounts are suitable for useful for a wide variety of samples such as fruit, vegetables and other solids that can be cut into small slices.

Smear

A smear is made by carefully smearing a thin layer of the specimen across a slide and then applying a cover slip. Typically, a smear should be allowed to air dry before applying a stain.

STAINS

Stains are used to help identify different types of cells using light microscopes. They give the image more contrast and allow cells to be classified according to their shape (morphology). By using a variety of different stains, you can selectively stain different areas such as a cell wall, nucleus, or the entire cell. Stains can also help differentiate between living or dead cells.

Stains tend to be grouped as neutral, acidic or basic, depending upon their chemical makeup and will attract or repel different organisms accordingly. For example, scientists and health professionals use Methylene Blue, a slightly alkaline stain, to reveal the presence of deoxyribonucleic acid, more commonly known as DNA.

Stain Types

Iodine is one of the more commonly available stains and is used to identify starch in a variety of samples. It will stain carbohydrates in plants and animal specimens brown or blue-black. Glycogen will show as red.

Methylene Blue is an alkaline stain useful in identifying acidic cell nuclei and DNA in animal, bacteria or blood samples. It&rsquos also useful in aquariums to prevent the spread of fungal infections in fish. See more details >

Eosin Y is an acidic stain which stains pink for alkaline cells (cytoplasm, for example). It colors red for blood cells, cytoplasm and cell membranes. Eosin's most important medical uses are in blood and bone-marrow testing, including the PAP smear. See more details >

Gram's Stain is one of the most frequently used processes in identifying bacteria &ndash used daily in hospitals. It is a primary test that quickly and cost effectively divides bacteria into one of two types: Gram positive or Gram negative. See more details >


Microscope Cover Slips

How do you know when to use a microscope cover slip? And if you do need to use one, what thickness is appropriate? This post will answer all of these questions and hopefully help you figure out what mediums will produce the best images under different microscopes.

What type of microscope are you using?

The first question you should answer is what type of microscope are you using? Some microscopes do not require the use of a cover slip at all. Below is a list of a variety of microscopes and their use of cover slips:

  • Stereo Microscopes - when using a stereo microscope you do not need to use a cover slip. The sample sits directly on the microscope stage and is not typically placed on a microscope slide at all.
  • Inverted Biological Microscopes - Petri dishes are used with inverted microscopes in order to contain living samples in liquid. Cover slips are not used with a Petri dish, but the thickness of the Petri dish can be important. We will talk more about this below.
  • Inverted Metallurgical Microscopes - when using an inverted metallurgical microscope the sample will be flat and may be polished. The sample is placed directly on the stage and no slide or cover slip is used.
  • Upright Biological Microscopes (Compound Microscopes) - Upright biological microscopes are sometimes referred to as compound microscopes. When using an upright biological microscope both a slide and cover slip is used.
  • Upright Metallurgical Microscopes - Upright metallurgical microscopes do not require the use of a slide or cover slip. Occasionally one is used if the sample is a powder and must be flattened or contained, but typically the sample is placed directly on the stage. Filter patches are also placed directly on the stage under an upright metallurgical microscope.
  • Polarizing Microscopes - polarizing microscopes may be used to view thin sections of rocks, minerals, or even powdery substances. Depending on the sample, a slide and cover slip is used at times to flatten and contain the sample.

When viewing sections or smears (typically biological in nature), they must be fixed to a slide. The sample is preserved or prepared with a medium (stain) by placing a cover slip on top of the sample on the slide. Thin sections of plant samples can be placed on a slide with a drop of water with a cover slip on top of the plant to flatten it.

Take a look at the microscope objective lens shown at left. This objective is a 20x plan achromat objective lens with a Numerical Aperture (NA) of 0.45. The infinity symbol tells us that it is an infinity corrected lens and after this symbol the lens shows "0.17". This refers to the cover slip thickness.

Standard transmitted light objective lenses are designed for a 0.17mm cover slip between the sample and the lens. Standard cover slips have a thickness of 0.13-0.16mm, taking into consideration the added layer of an embedding medium or water into account in the sample results in meeting the rule of 0.17mm.

The higher the Numerical Aperture of an objective lens, the higher the sensibility for deviations from the value on the objective lens. For example, a 4x/0.10 objective lens can be used with or without a cover slip and no difference will be noted. But if an objective lens with a value of 40x/0.95 were used, any variation off the cover slip requirement could result in an image that is not clear and crisp. In this situation using the appropriate cover slip thickness, as well as ensuring that samples were precisely cut in a thin section (1-5 microns thick) will result in the clearest high quality images.

Additionally, if too much of a solution is used between the cover slip and microscope slide (such as stain or liquid), it can ruin the image. When preparing slides it can help to press the cover slip into the slide with the tip of a needle (to avoid fingerprints on the slide while flattening it), as well as mopping up extra fluid with a paper towel placed at the edge of the cover slip.

Some objective lenses will be marked with a "0". This indicates that the objective lens does not require use of a cover slip at all. In some fields these objective lenses are used to save time by making slide preparation faster.

When using an inverted biological microscope, Petri dishes, flasks or well plates hold the microscopy samples. These vessels have a thickness of 1mm on the bottom. Therefore the objective lenses used with these vessels are marked with a 1.1mm indicator. The additional 0.1mm comes from the water or agar medium in the Petri dish.

Due to the improved working distance in inverted microscope objectives lenses, they are not driven to maximum resolution (NA). The inverted microscope was constructed for the improved handling freedom it provides with specimens and was not created for the evaluation of resolution limits. If you wish to use a 0.17mm objective lens on an inverted microscope, make sure to use a Petri dish with a 0.17mm glass bottom, or you could turn the glass slide upside down. Of course if you are using a slide containing liquid, turning it upside down will not be beneficial.

Next time you are preparing samples for your microscope, take a look at the objective lens and ensure you are using the proper cover slip thickness for the lens.

If you have questions regarding microscope cover slips and obtaining the best image with your objective lens, contact Microscope World and we will be happy to help.


3 - 1 Use of the microscope

The microscope, as shown in Figure 3-1, is one of the most important instruments utilized by the microbiologist. In order to study the morphological and staining characteristics of microorganisms such as bacteria, yeasts, molds, algae and protozoa, you must be able to use a microscope correctly.

Figure 3.1. The light microscope. A modern light microscope. This is an example of the kind used in the teaching labs at the University of Wisconsin-Madison. The various parts of the microscope are labeled. Please take the time to become familiar with their names.

The compound microscope used in microbiology is a precision instrument its mechanical parts, such as the calibrated mechanical stage and the adjustment knobs, are easily damaged, and all lenses, particularly the oil immersion objective, are delicate and expensive. Handle the instrument with care and keep it clean.

The microscope is basically an optical system (for magnification) and an illumination system (to make the specimen visible). To help understand the function of the various parts of the microscope, we will follow a ray of light as it works its way through a microscope from the light source, through the lenses, up to the eye. Figure 3-8 traces the path of light through the parts of the microscope

Figure 3.8. The path of light through a microscope. Modern microscopes are complex precision instruments. Light, originating in the light source (1), is focused by the condensor (2) onto the specimin (3). The light then enters the objective lens (4) and the image is magnified. Light then passes through a series of glass prisms and mirrors, eventually entering the eyepiece (5) where is it further magnified, finally reacing the eye.

First let us consider a primary feature of all microscopes, the light source. Proper illumination is essential for effective use of a microscope. A tungsten filament lamp usually serves as the source of illumination. If reflected illumination is used, a separate lamp provides a focused beam of light which is reflected upward through the condenser lenses by a mirror.

The light from the illuminating source is passed through the substage condenser. The condenser serves two purposes it regulates the amount of light reaching the specimen and it focuses the light coming from the light source. As the magnification of the objective lens increases, more light is needed. The iris diaphragm (located in the condenser), regulates the amount of light reaching the specimen. The condenser also collects the broad bundle of light produced by the light source and focuses it on the small area of the specimen that is under observation.

Light then passes up through the slide and into the objective lens where the first magnification of the image takes place. Magnification increases the apparent size of an object. In the compound light microscope two lenses, one near the stage called the objective lens and another in the eyepiece, enlarge the sample. The magnifying power of an objective lens is engraved in the lens mount. Microscopes in most microbiology laboratories have three objective lenses: the low power objective lens (10X), the high-dry objective lens (40X) and the oil-immersion objective lens (100X). The desired objective lens is rotated into working position by means of a revolving nosepiece.

On both sides of the base of the microscope are the course and fine adjustment knobs, used to bring the image into focus. Rotation of these knobs will either move the specimen and the objectives closer or farther apart. The coarse adjustment moves the nosepiece in large increments and brings the specimen into approximate focus. The fine adjustment moves the nosepiece more slowly for precise final focusing. In some microscopes, rotation of the fine and course adjustment knobs will move the stage instead of the nosepiece.

Magnification alone is not the only aim of a microscope. A given picture may be faithfully enlarged without showing any increase in detail. The true measure of a microscope is its resolving power. The resolving power of the lens is its ability to reveal fine detail and to make small objects clearly visible. It is measured in terms of the smallest distance between two points or lines where they are visible as separate entities instead of one blurred image. The resolving power of the objective lens, engraved on the lens, allows us to predict which objective lens should be used for observing a given specimen. However, having good resolution in the microscope does not guarantee a visible image, the resolving power of the human eye is quite limited. Often further magnification is needed to obtain a good image.

When the oil-immersion objective lens is in use, the difference between the light-bending ability (or refractive index of the medium holding the sample) and the objective lens becomes important. Because the refractive index of air is less than that of glass, light rays are bent or refracted as they pass from the microscope slide into the air, as shown in Figure 3-9. Many of these light rays are refracted at so great an angle that they completely miss the objective lens. This loss of light is so severe that images are significantly degraded. Placing a drop of immersion oil, which has a refractive index similar to glass, between the slide and the objective lens decreases this refraction, and increases the amount of light passing from the specimen into the objective lens. This results in greater resolution and a clearer image.

Figure 3.9. Refraction of light at 100X. Light passing out of the slide, into the air, toward the objective lens is refracted, due to the different in refractive index between air and glass. While the bending cause by this difference is not important at 100X and 400X, at 1000X this refraction is problematic, causing blurring of the image and significant loss of light. Immersion oil has a refractive index very similar to that of glass. Placement of a drop of oil between the objective lens and the slide prevents the bending of light rays and clarifies the image. The blue dashed line represents a potential light ray if immersion oil is not present. The red dashed line represents a light ray if immersion oil is present.

The image of the specimen continues on through a series of mirrors and/or prisms that bend it toward the eyepiece. A further magnification takes place at the eyepiece producing what is called a virtual image. Total magnification is equal to the product of the eyepiece magnification and the objective magnification. Most often eyepiece lenses magnify 10-fold resulting in total magnifications of 100, 400, or 1000X, depending upon which objective is in place. Many modern microscopes will also have focusable eyepieces to compensate for differences between individuals and even between individual's eyes. The adjustment of these is important and is described below.

3 - 2 Operating procedure

Below we describe detailed directions for the use of a microscope. This will give you an appreciation of their operation. These directions have been written as generally as possible, but it may be necessary for your instructor to make modifications for the exact microscopes you are using. Light microscopes used in teaching laboratories are designed for ease of use and with some practice should become automatic.

Raise the nosepiece using the course adjustment knob. This provides greater access for positioning the slide on the stage.

Rotate the nosepiece so that the 10X objective lens is in operating position.

Open the iris diaphragm approximately half way.

Turn on the in-base illuminator by depressing the push-type switch.

Place a stained specimen slide on the stage and with the naked eye position the specimen directly above the center of the condenser.

Use the thumbwheel below the eyepieces to adjust the interpupillary distance between the two eyepieces. This is important to be able to view specimens with both eyes, maximizing the quality of the image and preventing fatigue from prolonged use of one eye.

Move the microscope condenser by means of the condenser adjustment knob until the top of the condenser is almost at the highest position. There should be enough room to slide a piece of paper between the stage and the condenser, but no more. This will focus the light onto the slide.

Rotate the coarse adjustment knob in a clockwise direction to bring the 10X objective closer to the slide. View through the eyepieces and, without disturbing the coarse adjustment setting, slowly rotate the fine adjustment knob until the specimen is in the sharpest possible focus.

The left eyepiece tube is focusable to compensate for refraction differences of the eyes. The correct procedure is to bring the specimen into sharp focus looking though the right eyepiece only. Then focus for the left eye by turning the left eye tube collar fully counter-clockwise. Next, while viewing the specimen with the left eye only turn the knurled collar clockwise until the specimen is in sharp focus. Do not adjust the fine adjustment knob during this procedure.

Remove an eyepiece to view the back aperture of the objective lens. Close the condenser iris diaphragm, then re-open until the leaves of the diaphragm just disappear from view. Replace the eyepiece and view the specimen. The iris diaphragm may be closed slightly to enhance contrast, especially when viewing unstained specimens.

Unstained specimens have only minimal contrast with their surrounding environments. As a result they can usually be viewed more effectively by setting the diaphragm at or near minimum opening. Reducing the diaphragm setting increases definition, contrast, and depth of focus but introduces diffraction problems and sacrifices resolution. Play with the diaphragm setting and select the best compromise by trial and error.

Once the specimen is in sharp focus using the 10X objective lens, it is then possible to rotate the nosepiece to the 40X objective lens without changing the position of the coarse adjustment knob. Very little refocusing with the fine adjustment knob is required since most light microscope objective lenses are parfocal. Remember that the iris diaphragm setting must be changed to allow more light to pass though the sample as the magnification increases.

If the specimen is to be viewed using the 100X oil immersion lens, immersion oil must be applied to the slide.

Rotate the 40X objective lens slightly to the side so that a drop of immersion oil may be placed on the specimen without getting it on the 40X lens.

Place a drop of immersion oil in the center of the circle of light formed on the specimen slide.

Carefully turn the nosepiece until the 100X objective lens snaps into place. The objective lens should be in the oil but must not touch the slide.

Increase the light intensity as required and rotate the fine adjustment knob to obtain a sharp focus of the specimen. If necessary make further adjustments to obtain optimal illumination.

If the microscope is not parfocal, it will be necessary to lower the objective lens as close to the slide as possible without touching it. This is done only while looking at the lens and slide from the side of the microscope. Bring the specimen into view by slowly raising the objective lens with the coarse adjustment knob. Next, focus with the fine adjustment knob and adjust the illumination as necessary. If this is not successful the first time, repeat the entire procedure.

In many cases, a preparation needs to be observed only under the oil immersion lens. In this case, first locate the specimen and center it in the field with the low power objective lens. Then add oil and rotate the oil immersion objective lens into position.

3 - 3 Common Problems

Certain problems real or apparent, may be encountered while operating your microscope. Here is a trouble shooting guide to help you if you are having difficulty focusing a sample.

The sample can be focused at 10X, but it is difficult to find or blurry at 40X.

This is often caused by immersion oil on the 40X lens. Wipe the 40X lens with lens paper to remove the oil and refocus. This can be prevented by never viewing a specimen with the 40X objective after adding immersion oil to a slide.

The sample can be focused at 10X but when the 40X lens is rotated in place it contacts the slide.

In most cases this is caused by the slide being place on the stage upside down †with the smear facing the stage. Check your slide carefully to make sure it is placed on the stage correctly.

The fine adjustment knob does not turn in the direction required for sharp focusing.

This indicates that it has been turned to the limits of its threads, either upward or downward, as the case may be. Screw it back to about one-half the thread distance (About four turns), use the coarse adjustment to raise or lower the objective lens sufficiently to bring the specimen into view then refocus with the fine adjustment.

What I am viewing does not look like bacteria.

Check to make sure you are in the correct focal plane that you are focusing on the smear and not dust on the lenses. To verify this, move the slide while looking at it. Anything in the smear should move in the field of view.

What I am viewing does not look like bacteria. Part II

If you are in the correct focal plane, there may be problems with smear preparation. Did you heat fix too much? Was the amount of culture applied sufficient? Did you stain the slide correctly? Many apparent microscope problems can be attributed to poor slide preparation.

3 - 4 Proper care of the microscope

The following rules, cautions and maintenance hints will help keep your microscope in good operating condition.

Use both hands when carrying the microscope: one firmly grasping the arm of the microscope the other beneath the base. Avoid jarring your microscope.

To keep the microscope and lens systems clean:

Never touch the lenses. If the lenses become dirty, wipe them gently with lens tissue.

If blurred specks appear in the field of view this may be due to lint or smears on the eyepiece. If the specks move while rotating the eyepiece, the dust is on the eyepiece and cleaning the outer lens of the eyepiece is in order. If the quality of the image is improved by changing objective lenses, clean the objective lens with lens paper.

Never leave a slide on the microscope when it is not in use.

Always remove oil from the oil-immersion objective lens after its use. If by accident oil should get on either of the lower-power objective lenses, wipe it off immediately with lens tissue.

Keep the stage of the microscope clean and dry. If any liquids are spilled, dry the stage with a piece of cheesecloth. If oil should get on the stage moisten a piece of cheesecloth with xylol and clean the stage, then wipe it dry.

When not in use, store your microscope in its cabinet. Put the low power objective lens into position at its lowest point above the stage. Be sure that the mechanical stage does not extend beyond the edge of the microscope stage. Wrap the electrical cord around the base.

To avoid breaking the microscope:

Never force the adjustments. All adjustments should work freely and easily. If anything does not work correctly, do not attempt to fix it yourself, immediately notify your instructor.

Never allow an objective lens to jam into or even to touch the slide or cover-slip.

Never focus downward with the coarse adjustment while you are looking through the microscope. Always incline your head to the side with eyes parallel to the slide and watch the objective as you move it closer to the slide. This will prevent you from smashing the objective into the slide.

Never exchange the objective or eyepiece lenses of different microscopes, and never under any circumstances remove the front lenses from objective lenses.

Never attempt to carry two microscopes at one time

If you follow these rules, you will never have trouble with your microscope.

3 - 5 Staining microorganisms

Preliminary identification of bacteria is usually based upon their cell morphology and grouping and the manner in which they react to certain staining procedures. The purpose of this section is to demonstrate some common staining reactions used to categorize microorganisms.

Figure 3.2. An unstained bacterial smear. Unstained bacteria are mostly made of water and are nearly transparent when viewed through a light microscope (pictured on the left). Note that most of the microbes are not visible, but a dust spec in the center of the field of view is visible. Stains cling to the positive and negative charges of bacteria, but do not bind as readily to the background of a slide. They therefore differentiate microbes from their surroundings. Stained bacteria are shown at 40X and 100X in the center and right panels.

Unstained bacteria are practically transparent when viewed using the light microscope and thus are difficult to see as shown in Figure 3-2. The development of dyes to stain microorganisms was a significant advance in microbiology. Stains serve several purposes:

Stains differentiate microorganisms from their surrounding environment

They allow detailed observation of microbial structures at high magnification

Certain staining protocols can help to differentiate between different types of microorganisms.

Most dyes consist of two functional chemical groups as shown in Figure 3-3. The chromophore group, which give dyes their characteristic color and the auxochrome group, containing an ionizable chemical structure, which helps to solubilize the dye and facilitates binding to different parts of microorganisms. Previously, dyes were classified as acidic or basic, depending upon whether the pigment was negatively or positively charged at neutral pH. More accurately, dyes can be referred to as anionic (-) or cationic (+) and this is the convention that will be used in this manual. Cationic dyes (crystal violet, methylene blue) will react with groups on bacteria that have a negative charge. Anionic dyes (eosin, nigrosine) will react with groups that have a positive charge. Since most bacteria have many positive and negative groups in their cell walls and other surfaces, they will react with both cationic and anionic dyes.

Figure 3.3. The structure of crystal violet. The auxochrome groups of crystal violet is the charged carbon in the center of the molecule. This is typically neutralized by a Cl - ion. The chromophore group consists of the three benzene rings and the central carbon. These structures readily absorb light.

Staining protocols can be divided into 3 basic types, simple, differential, and specialized. Simple stains react uniformly with all microorganisms and only distinguish the organisms from their surroundings. Differential stains discriminate between various bacteria, depending upon the chemical or physical composition of the microorganism. The Gram stain is an example of a differential stain. Specialized stains detect specific structures of cells such as flagella and endospores.

3 - 6 Preparation of a Bacterial Smear for Staining

Before staining and observing a microbe under a microscope, a smear must be prepared. The goal of smear preparation is to place an appropriate concentration of cells on a slide and then cement them there so that they do not wash off during the subsequent staining procedure. Figure 3-4 demonstrates smear preparation.

The best smears are made from bacteria that have grown on a solid surface such as an agar slant or plate. A bit of growth from a culture is mixed with distilled or tap water to form a slightly turbid solution and this is spread on a clean grease free slide. When staining broth cultures, a drop of broth is transferred directly to a slide, using no extra water. The procedure for making a smear is as follows:

If more than one culture is to be examined using the same stain, it is possible to prepare up to 6 smears on the same slide. Before preparing the slide, divide it into the appropriate number of sections and clearly label each section on the underside of the slide.

If your culture has been grown on a agar slant or agar plate. Place a small drop of water on a clean, grease-free slide. Next, using a sterile loop or straight wire needle, transfer a bit of the growth to the drop of water and rub the needle around until the material is evenly emulsified. Spread the drop over a portion of the slide to make a thin film. The suspension should be only slightly turbid.

If you are using a broth culture, the broth culture must have clearly visible turbidity. Transfer a loopful of culture from the broth onto a clean grease free slide. Spread the drop over a portion of the slide to make a thin film.

Allow the film to air-dry. To get a good stain, it is important to let the smear dry completely. Excess water left on the slide will boil during the fixing stage, causing most microbe present to rupture. Rushing this step will result in a poor final stain.

Once dry, "fix" the smear to the slide by passing the bottom of the slide through the tip of the burner flame several times for a one second. After heat fixing, touch the heated portion of the slide to your hand. It should be comfortably warm, but not burning hot.

Take care not to under-fix (the smear will wash off) or over-heat (the cells will be ruptured or distorted) the slide. The correct amount of heat fixing is learned by experience.

Allow the smear to cool and apply the stain.

3 - 7 The Simple Stain

In a simple stain, the smear is stained with a solution of a single dye which stains all cells the same color. Differentiation of cell types or structures is not the objective of the simple stain. However, certain structures which are not stained by this method may be easily seen, for example, endospores and lipid inclusions.

Simple stains are, well simple. One makes a smear and the applies a single stain to the slide. Below is a procedure for a simple stain.

Prepare and heat-fix a smear of the organism to be studied.

Cover the smear with the staining solution. If crystal violet or safranin is used, allow one minute for staining. The use of methylene blue requires 3-5 minutes to achieve good staining.

Carefully wash off the dye with tap water and blot the slide dry with blotting paper, an absorbent paper pad or a paper towel.

Three steps, now wasn't that easy?

The above movie demonstrates the simple stain.

Figure 3-10 shows a light micrograph of what a simple stain should look like.

Figure 3.10. The Simple Stain. A photomicrograph of a simple stain at 1000X magnification. Note that all cells, regardless of species or cell wall construction, stain the same color.

3 - 8 The Gram Stain

The Gram stain, performed properly, differentiates nearly all bacteria into two major groups. For example, one group, the gram-positive bacteria, include the causative agents of the diseases diphtheria, anthrax, tetanus, scarlet fever, and certain forms of pneumonia and tonsillitis. A second group, the gram-negative bacteria, includes organisms which cause typhoid fever, dysentery, gonorrhea and whooping cough. In Bacteria the reaction to Gram stain reagents is explained by different cell wall structures. Gram-positive microbes have a much thicker cell wall, while that found in Gram-negative microbes is thinner. Microbes from the Archaea domain contain different cell wall structures than that seen in microbes commonly found in the lab (Bacteria domain). However, they will still have a species specific Gram stain reaction, even though the underlying macromolecular structures are different.

The Gram stain is one of the most useful differential stains in bacteriology, including diagnostic medical bacteriology. The differential staining effect correlates to differences in the cell wall structure of microorganisms (at least Bacteria, but not Archaea as mentioned above). In order to obtain reliable results it is important to take the following precautions:

The cultures to be stained should be young - incubated in broth or on a solid medium until growth is just visible (no more than 12 to 18 hours old if possible). Old cultures of some gram-positive bacteria will appear Gram negative. This is especially true for endospore-forming bacteria, such as species from the genus Bacillus. In this class, many of the cultures will have grown for more than 2 days. For most bacteria this is not a problem, but be aware that some cultures staining characteristics may change!

When feasible, the cultures to be stained should be grown on a sugar-free medium. Many organisms produce substantial amounts of capsular or slime material in the presence of certain carbohydrates. This may interfere with decolorization, and certain Gram-negative organisms such as Klebsiella may appear as a mixture of pink and purple cells.

Gram stain procedure

Below is a procedure that works well in the teaching laboratories.

Cover the slide with crystal violet stain and wait one minute.

After one minute wash the stain off (gently!) with a minimum amount of tap water. Drain off most of the water and proceed to the next step. It may help to hold the slide vertically and touch a bottom corner to paper toweling or blotting paper.

Cover the slide with iodine solution for one minute. The iodine acts as a mordant (fixer) and will form a complex with the crystal violet, fixing it into the cell.

Rinse briefly with tap water.

Tilt the slide lengthwise over the sink and apply the alcohol-acetone decolorizing solution (dropwise) such that the solution washes over the entire slide from one end to the other. All smears on the slide are to be treated thoroughly and equally in this procedure. Process the sample in this manner for about 2-5 seconds and immediately rinse with tap water. This procedure will decolorize cells with a Gram negative type of cell wall but not those with a gram-positive type of cell wall, as a general rule. Drain off most of the water and proceed.

As the decolorized gram-negative cells need to be stained in order to be visible, cover the slide with the safranin counterstain for 30 seconds to one minute.

Rinse briefly and blot the slide dry. Record each culture as Gram positive (purple cells) or Gram negative (pink cells).

The above video demonstrates the Gram stain procedure, while Figure 3-11 shows the results of a Gram stain for gram-positive and gram-negative negative bacteria.

Figure 3.11. The Gram Stain. A photomicrograph of gram-positive and gram-negative bacteria. Note that Gram reaction is dependent upon cell wall structure. A) E. coli a common gram-negative rod found in the colon. B) Staphylococcus epidermidis a gram-positive cocci found on the skin. C) Bacillus cereus a gram-positive rod found in the soil.

3 - 9 The Endospore Stain

Cells of Bacillus, Desulfotomaculum and Clostridium (and several other, lesser-known genera--see Bergey's Manual) may, as a response to nutrient limitations, develop endospores that possess remarkable resistance to heat, dryness, irradiation and many chemical agents. Each cell can produce only one endospore. It is therefore not a reproductive spore as seen for some organisms such as Streptomyces and most molds. The endospore is essentially a specialized cell, containing a full complement of DNA and many proteins, but little water. This dehydration contributes to the spores resistance and makes it metabolically inert. The endospore develops in a characteristic position (for its species) in the vegetative cell. Eventually the cell lyses, releasing a free endospore.

Endospore Stain Procedure

Endospore stains require heat to drive the stain into the cells. For a endospore stain to be successful, the temperature of the stain must be near boiling and the stain cannot dry out. Most failed endospore stains occur because the stain was allowed to completely evaporate during the procedure.

Place the heat-fixed slide over a steaming water bath and place a piece of blotting paper over the area of the smear. The blotting paper should completely cover the smear, but should not stick out past the edges of the slide. If it sticks out over the edges stain will flow over the edge of the slide by capillary action and make a mess.

Saturate the blotting paper with the 5-6% solution of malachite green. Allow the steam to heat the slide for five minutes, and replenish the stain if it appears to be drying out.

Cool the slide to room temperature. Rinse thoroughly and carefully with tap water.

Apply safranin for one minute. Rinse thoroughly but briefly with tap water, blot dry and examine. Mature endospores stain green whether free or in the vegetative cell. Vegetative cells stain pink to red.

The above video demonstrates the endospore stain

Figure 3-12 shows a photomicrograph of an endospore stain.

Figure 3.12. The Endospore Stain. A photomicrograph of an enodspore stain. Spores present in the picture stain green, while the vegetative cells stain red. A) Staphylococcus epdiermidis which does not form endospores. B) The endospore-forming rod, Bacillus cereus.

3 - 11 Practice staining

In the study and identification of bacteria, the microscope is indispensable! The series of micro-scopic observations in this exercise is designed to illustrate how bacteria may be viewed individually in their basic form, the cell. The second and third periods herein coincide with those of Experiment 1 where organisms isolated by the student are examined microscopically (and could be found to be more interesting than those provided in this exercise!).

Period 1

Materials

Hay infusions and various other items from nature

Slide with smears of Bacillus cereus and Staphylococcus epidermidis

Simple stain.

  1. You are provided with a microscope slide with two smears. Following the directions for microscopy and staining, heat-fix the slide, making sure the slide goes through the flame smear-side up.
  2. Gloves are available for the staining procedure. Placing the slide on the staining rack in the sink, cover the slide with crystal violet for one minute. For a review, look at the directions for the simple stain.
  3. Carefully rinse off the dye with tap water and blot the slide dry with paper towel or blotting paper.
  4. With both hands, obtain the light microscope from the cabinet (corresponding to your desk number). This is the type of microscope which we will always use to observe stained smears.
  5. Unless the instructor has other directions more directly applicable to the microscope you are using, use the simple procedure described in the operating procedure. Refer to this procedure as you study the cells in the two smears in the following steps.
  6. Place the slide on the stage such that it is oriented as illustrated above. Make sure the clips on the stage hold the slide securely.
  7. Begin your observations with the Bacillus cereus smear. (See figure below) When observing this organism with the oil-immersion objective, you will notice that the cells are relatively large and rod-shaped (bacilli) and are usually in chains. Record your observations on the next page.
  8. Repeat this procedure with the Staphylococcus epidermidis smear. Cells of this organism are spheres (cocci) which are usually arranged in clusters (staphylococci) and pairs.
  9. When you are through, be sure the microscope is put away properly (i.e., all oil wiped off, 10X objective lens in place, stage centered). It is recommended that you keep the slide. (To remove immersion oil from smears, place a few pieces of lens paper on the slide to absorb the oil. Then, add several drops of xylol to the lens paper. Peel the paper, now soaked with xylol, off the slide. Xylol is flammable! Keep it away from flames!)

Figure 3.13. Simple stain. A simple stain of S. epidermidis and B. cereus. S. epidermidis (A), B. cereus old (B), B. cereus young (C)

Wet Mount

  1. For observation of living microorganisms, various samples including a hay infusion are available. To study the microorganisms in the aqueous materials available, it is necessary to make wet mounts. The procedure is relatively simple:
  2. Using a capillary pipette or inoculating loop, pick up some of the material from around the surfaces of grass and leaves and from the bottom of the sample. Place a drop of suspended material on a clean microscope slide.
  3. With a toothpick, spread a very thin layer of vaseline over a small part of the palm of your hand. Take a clean coverslip (always held by the edges) and gently scrape all four edges along your palm, picking up a thin line of vaseline along each edge.
  4. Place the coverslip directly onto the drop on the slide in such a manner that some air bubbles are trapped. Place a small, multilayered piece of paper towel over the coverslip and press down. Discard the piece of paper towel into the disinfectant.
  5. Examine the wet mount with your light microscope or a phase CONTRAST microscope set up by the instructor at a special station in the back or side of the lab.
  6. Without removing the coverslip, discard the slide into the disinfectant container on the stage. (Refer to page viii for cleanup directions.)

If you haven't already, Figure 1-2 presents a movie of the types of life forms found in a hay infusion.

Period 2

Materials

Bacterial cultures growing either in a liquid medium (Heart Infusion Broth) or on a slant of an all-purpose medium followed by suspension in saline:

Escherichia coli - young culture, incubated 12-15 hours

Bacillus cereus - young culture, incubated 12-15 hours

Bacillus cereus - old culture, incubated 2-3 days

Figure 3.14. Gram stains. Gram stains of demonstration species. Below are shown typical Gram stain reactions of two species. E. coli (A), B. cereus old (B), B. cereus young (C). The images are slightly larger than what would be visible in a light microscope to improve clarity.

  1. On one clean glass slide, prepare smears of the three cultures. Go to smear prepapration if you need a refresher. Only when the smears have dried completely should the slide be heat-fixed.
  2. Perform the Gram stain procedure as described.
  3. As with any stained smear, definitive observations are made with the 100X, oil-immersion objective. Refer to the microscope directions already given, remembering to focus the slide initially with the 10X objective, moving then to the oil immersion objective.
  4. Keep in mind that the young cultures of B. cereus and E. coli are your positive and negative control cultures, respectively, for the observation of probable gram-variability of the older B. cereus culture.
  5. Using the figures below, record your observations in your notebook, noting the Gram reaction (positive if purple, negative if red) and the cellular shape. Is there any difference seen between the two cultures of Bacillus cereus? Is gram-variability evident for the older culture? Recall from the introduction to Experiment 1 that we can refer to old and young cultures but should not do so for individual cells. (Remember to discard the tubes and slides properly)

Period 3

Materials

This experiment will be done in class.

Young bacterial cultures growing on slants of Heart Infusion Agar:

Staphylococcus epidermidis

Pseudomonas fluorescens

Record the number of your unknown!

Figure 3.15. Typical reactions of example strains for test. The classic Gram reactions for Staphylococcus epidermidis (A) and Pseudomonas fluorescens (B). From this, determine whether they are Gram (+) or Gram (-). Note we do not show an unknown as this must be done in class. The images are slightly larger than what would be visible in a light microscope to improve clarity.

  1. On a clean glass slide, prepare heat-fixed smears of the three cultures, noting that these cultures are growing on a solid medium. Therefore the cells must be dispersed in a drop of water when preparing the smears, as a smear is always a dried suspension of cells. Take care not to make the smears too thick! S. epidermidis and P. fluorescens are your positive and negative control cultures (respectively) for your unknown.
  2. Perform the Gram stain procedure and note the Gram reaction and cellular shape. Record your results. Fill out and turn in your description of your unkonwn. Save your slide until your graded unknown is returned.

Period 4

Materials

36-48 hour culture of Klebsiella pneumoniae growing on a slant of EMB Agar (a high-sugar medium)

Dropper bottle of filtered India ink

3-day culture of Mycobacterium smegmatis growing on a slant of Trypticase Soy Agar plus 1% glycerol

18-24 hour culture of Micrococcus luteus (the negative control culture) growing in Nutrient Broth

Dropper bottles of carbol fuchsin (freshly-made), acid alcohol and methylene blue

Capsule stain

Figure 3.16. The capsule stain. A capsule stain using India ink at 1000x magnification. The cells of Klebsiella pneumoniaeare surrounded by a dark background. The capsule is the clear area surrounding the cells. The photomicrographs is slightly enlarged for clarity.

  1. Using the culture of Klebsiella pneumoniae, Place one loopful of water on a slide and emulsify in it a bit of growth from the slant or plate culture of the designated organism. Add a drop of filtered India ink to the cell suspension. It often works out well to place the drop of India ink adjacent to the cell suspension on the glass slide.
  2. Obtain a clean coverslip (no fingerprints, smudges, dirt, etc.) and rim it lightly with vaseline the vaseline can be gently scraped from a thin layer applied to the palm of the hand. Place a small, multi-layered piece (about 1-2 cm2) of paper towel over the coverslip and press down firmly discard the paper towel into the disinfectant.
  3. Using the regular light microscope, focus initially with the 10X objective, switching to the 45X objective and then - if needed - the 100X, oil-immersion objective. Adjust the light intensity as required with the iris diaphragm. The outline of the cell can be seen within the area of the clear capsule.
  4. Alternatively, the phase microscope can be used. Heed the precautions regarding use of this microscope. Excellent observations can be made with just the 40X objective lens (which takes no immersion oil).
  5. When finished, without removing the coverslip, discard the slide directly into the disinfectant. Never discard capsule stains and other wet mounts with the stained smears, as viable cells are still present and the slides must be disinfected!. Record your observations below.
Acid-fast stain

Figure 3.17. The acid fast stain. A photomicrograph of Mycobacterium smegmatis (pink) and Micrococcus luteus (blue) at 1000x magnification. M. smegmatis is acid-fast, retaining the carbol fuchsin dye, thus appearing pink. M. luteus is not acid-fast, loses the carbol fuchsin during decolorizaiton, and is counter-stained with methylene blue.

  1. Prepare a mixed smear of two organisms as follows: Place a drop of the Micrococcus luteus broth culture on a slide. Into this drop, add cells from the Mycobacterium smegmatis culture. Disperse the cells as much as you can (the Mycobacterium cells tend to clump), and prepare a smear about the size of a nickel. Let it air-dry completely, and then heat-fix it well, passing the slide through the flame an extra one or two times.
  2. Perform the acid-fast procedure (page 148, observing the slide with the regular light microscope) and record your observations below.
  3. As with all stained smears, discard the slide in the appropriate container.

3 - 12 Summary of Microscopy and Staining

Staining and viewing microbes under the microscope is often necessary in for their identification and classification. The identity of a microbe can help in determining the cause of a disease or the source of food spoilage. Microscopes also have important roles in genetics, cell structure, biochemistry and many other scientific disiplines. Hopefully, this short introduction has helped you to understand the visualization of microorganisms.


Types of light microscopes

The bright field microscope is best known to students and is most likely to be found in a classroom. Better equipped classrooms and labs may have dark field and/or phase contrast optics. Differential interference contrast, Nomarski, Hoffman modulation contrast and variations produce considerable depth of resolution and a three dimensional effect. Fluorescence and confocal microscopes are specialized instruments, used for research, clinical, and industrial applications.

Other than the compound microscope, a simpler instrument for low magnification use may also be found in the laboratory. The stereo microscope, or dissecting microscope usually has a binocular eyepiece tube, a long working distance, and a range of magnifications typically from 5x to 35 or 40x. Some instruments supply lenses for higher magnifications, but there is no improvement in resolution. Such "false magnification" is rarely worth the expense.


Ideas for discussing the demonstrations in class

Water molecules have a strong mutual attraction for one another, enabling water molecules to hold together strongly. Water and glass have a smaller attractive force, called adhesion rather than cohesion to distinguish the intermolecular forces between two separate bodies (glass attracting water) from those of water attracting water. Adhesion between water and glass leads to capillary rise in a glass tube. The cohesive force of water molecules is responsible for the phenomenon of surface tension.


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