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I'm posting this as a follow-up on What to look for when buying a light microscope?. The answerer states that you would need to use a an inverted microscope to count cell in the hemocytometer(counting chamber).
I've read, partly, a book on lab diagnostics tests and it was not written that there. Moreover, they mentioned things about microscopes featuring a compound upright microscope both in words and picture.
Yes it can. You will want to use a thin hemocytometer and the working distance of the objective (20x) needs to be long enough to focus both the cells under the cover glass and the grid under the hemocytometer. Because you are looking at it from the top it would be helpful probably to have some phase on the scope to increase your contrast so that you can more clearly see the grid.
Hemocytometers are most commonly used with inverted microscopes because inverted microscopes are the most convenient microscope for performing tissue culture, and 's a commonly used to count cells when passaging or setting up an experiment etc, as they allow you to view cells in a culture plate. Upright microscopes require that you make a slide.
You can use a hemocytometer(counting chamber) on whatever microscope for transmitted light, regular scope, inverted, whatever. It doesn't make any difference.
But you should keep in mind what a counting chamber is designed for: to count red or white blood cells, reticulocytes, yeast cells, whatever.
They're not designed with having the best possible image in mind. Their dimensions fall far outside the specifications microscope optics are calculated for.
I measured a few (regular Thoma from the well known brand Assistent): thickness is 4 mm. The standardized coverslips to be used with those are 0.4 mm thick.
There's a very small chance, some minor distortion will be visible in the image ;-)…
Brightfield microscopy is the most elementary form of microscope illumination techniques and is generally used with compound microscopes.
The name "brightfield" is derived from the fact that the specimen is dark and contrasted by the surrounding bright viewing field. Simple light microscopes are sometimes referred to as brightfield microscopes.
How it Works
In brightfield microscopy a specimen is placed on the stage of the microscope and incandescent light from the microscope’s light source is aimed at a lens beneath the specimen. This lens is called a condenser.
Featured right: Algae under the microscope with visible cells using brightfield illumination.
The condenser usually contains an aperture diaphragm to control and focus light on the specimen light passes through the specimen and then is collected by an objective lens situated in a turret above the stage.
The objective magnifies the light and transmits it to an oracular lens or eyepiece and into the user’s eyes. Some of the light is absorbed by stains, pigmentation, or dense areas of the sample and this contrast allows you to see the specimen.
For good results with this microscopic technique, the microscope should have a light source that can provide intense illumination necessary at high magnifications and lower light levels for lower magnifications.
Uses and Advancements
To some extent, brightfield microscopy is used in most disciplines requiring microscopic investigation.
Because it is a simple method, this is the first type of microscopy students learn in schools.
The life sciences, particularly microbiology and bacteriology, have always relied on the brightfield technique.
This technique can be used to view fixed specimens or live cells. Since many organic specimens are transparent or opaque, staining is required to cause the contrast that allows them to be visible under the microscope.
Different stains and staining techniques are used depending upon the type of specimen and cell structure being examined.
- Fuchsin is used to stain smooth muscle cells
- Methylene blue is used to stain cell nuclei is used on bacteria and gives rise to the name gram-negative or gram-positive bacteria based on the reaction of the bacteria to the stain. In fact, many scientific journals will not accept microbiological research for publication that is not supported by gram staining and brightfield illumination methodology. Most routine medical microscopic examination of blood and tissue is performed using this illumination technique.
Different complimentary techniques can be used to augment brightfield microscopy. By using a polarizing filter this illumination technique can be used in geological microscopic research and will reveal details not visible using white light.
Properly stained, microorganisms may be magnified to 1200x utilizing an oil immersion objective will increase resolution at this high magnification.
Digital Imaging Options
Although a basic method of microscopy, brightfield as a technique is well suited to mating with new technologies.
Digital imaging systems can make high resolution images of properly stained microorganisms using this technique.
Three-dimensional imaging accessories can be used with the brightfield method and newer technologies will allow real time viewing in 3D.
Also suited to video imaging, this enhancement will allow the user to view motile organisms interacting with their environment.
Brightfield technique has been mated with cell imaging software to better perform tasks previously delegated to fluorescence microscopy . By using multiple focal levels the cell borders and nuclei can be located in cell populations.
The benefit of using brightfield illumination for this task is that it frees fluorescent channels in microscopes and eliminates distortions caused by the overlapping of the color emissions of the stains and the excitation of the fluorescing materials.
Brightfield microscopy is very simple to use with fewer adjustments needed to be made to view specimens.
Some specimens can be viewed without staining and the optics used in the brightfield technique don’t alter the color of the specimen.
It is adaptable with new technology and optional pieces of equipment can be implemented with brightfield illumination to give versatility in the tasks it can perform.
Certain disadvantages are inherent in any optical imaging technique.
- By using an aperture diaphragm for contrast, past a certain point, greater contrast adds distortion. However, employing an iris diaphragm will help compensate for this problem.
- Brightfield microscopy can’t be used to observe living specimens of bacteria, although when using fixed specimens, bacteria have an optimum viewing magnification of 1000x.
Brightfield microscopy has very low contrast and most cells absolutely have to be stained to be seen staining may introduce extraneous details into the specimen that should not be present.
Also, the user will need to be knowledgeable in proper staining techniques.
Lastly, this method requires a strong light source for high magnification applications and intense lighting can produce heat that will damage specimens or kill living microorganisms.
Identify the ocular micrometer. A typical scale consists of 50 - 100 divisions. You may have to adjust the focus of your eyepiece in order to make the scale as sharp as possible. If you do that, also adjust the other eyepiece to match the focus. Any ocular scale must be calibrated, using a device called a stage micrometer. A stage micrometer is simply a microscope slide with a scale etched on the surface. A typical micrometer scale is 2 mm long and at least part of it should be etched with divisions of 0.01 mm (10 µm).
Suppose that a stage micrometer scale has divisions that are equal to 0.1 mm, which is 100 micrometers (µm). Suppose that the scale is lined up with the ocular scale, and at 100x it is observed that each micrometer division covers the same distance as 10 ocular divisions. Then one ocular division (smallest increment on the scale) = 10 µm at 100 power. The conversion to other magnifications is accomplished by factoring in the difference in magnification. In the example, the calibration would be 25 µm at 40x, 2.5 µm at 400x, and 1 µm at 1000x.
Some stage micrometers are finely divided only at one end. These are particularly useful for determining the diameter of a microscope field. One of the larger divisions is positioned at one edge of the field of view, so that the fine part of the scale ovelaps the opposite side. The field diameter can then be determined to the maximum available precision.
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.
Can one use a hemocytometer on a compound upright microscope? - 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.
Why Do Compound Microscopes Invert the Images?
The reason compound microscopes invert images lies in the focal length of the objective lens. The image focused by the lens crosses before the eyepiece further magnifies what the observer sees, and the objective lens inverts the image because of the lens' curvature. Digital microscopes that project images onto a screen correct for this problem, but laboratory-grade compound microscopes invert images, meaning they are upside down to the observer.
The inverted image is made from a positive lens, which means the image formed after light passes through the lens is a real image. This real image is inverted at the focal length. An example of this is using a letter of the alphabet. When the letter "e" is put right-side up in the slide to the observer, it is projected upside down in the tube. Moving the slide to the right shifts the image to the left, and vice versa.
A compound microscope is so called because there are multiple lenses magnifying images. Underneath the slide is a light source, then the stage upon which the slide sits. The image is refracted through the objective lens, and it travels up the body tube where the ocular lens magnifies the image a little more. The objective lens is where most of the magnification occurs, and many microscopes have rotating lenses that increase magnifications.
Compound Microscope – Types, Parts, Diagram, Functions and Uses
A compound microscope is a laboratory instrument used to magnify the image of a small object usually objects that cannot be seen by the naked eye.
It comes with two or more lenses, which causes it to achieve a higher level of magnification when compared with other low power microscopes. A compound microscope has the following:
- It comes with two or more convex lenses.
- One objective is used at a time.
- It produces 2-dimensional images.
- Its typical magnification is between 40x and 1000x.
- It is available in different configurations: monocular, binocular, and trinocular. (1, 2, 3, and 4)
Image 1: The image is a typical compound microscope commonly found in the workplace.
Who invented the compound microscope?
The invention of the compound microscope is credited by historians to Zacharias Janssen, a Dutch spectacle maker, around 1590.
Principles of compound microscope
When a minute object is placed beyond the focus of the objective lens, a highly magnified object is formed at a distance of distinct vision from the eye close to the eye piece. A compound microscope has two convex lenses an objective lens and eye piece.
The objective lens is placed towards the object and the eyepiece is the lens towards our eye. Both eyepiece and objective lenses have a short focal length and fitted at the free ends of two sliding tubes. (4, 5, and 6)
Compound microscope parts and magnification
A compound microscope consists of different parts and each part plays an important function. These include the following:
Image 2: The eyepiece/ocular lens of a compound microscope.
- Eyepiece/ocular lens – It is the part of the microscope that is looked through at the top. It comes with a magnification ranging between 5x and 30x.
Image 3: The head connects the eyepiece to the objective lens.
- Head (monocular/binocular) – It is the structural support of the microscope. It holds and connects the eyepiece to the objective lens.
Image 4: The objectives of a compound microscope.
- Objective lens – A compound microscope has three to five optical lens objectives and each comes with various magnification level (4x, 10x, 40x, and 100x). To calculate the total magnification of the microscope, all you need to do is to multiply the objective lens magnification by eyepiece magnification level.
Image 5: The arm of the compound microscope.
Image 6: The revolving nosepiece.
- Nosepiece – It holds the objective lens and attaches them to the head of the microscope. You can rotate the nosepiece to change the objective lens.
Image 7: The base is the bottom part of the microscope.
- Base – It supports the microscope and houses the illumination of the microscope.
Image 8: A sliding glass is needed and is attached to the stage using a stage clip.
- Sliding glass – It holds the specimen for easy viewing. It is made of thin rectangular glass.
Image 9: This is how a stage clip looks like.
Image 10: The stage is where the sliding glass with a specimen is placed,
Image 11: The aperture diaphragm control.
- Aperture – It is disc characterized by its circular opening where the illumination from the base reaches the platform stage.
Image 12: The condenser is underneath the stage.
- Abbe condenser – It is a lens that condenses the light from the base illumination and directed it to the stage.
Image 13: The coarse and fine adjustment knobs.
- Adjustment controls/knob (coarse/fine) – It allows you to easily adjust the focus of the microscope. By adjusting the knob, you can easily increase or decrease the level of details seen when examining at the slide through the eyepiece.
- Coarse focus – Use this knob with the lowest power objective to get the subject in focus.
- Fine focus – It is the smaller of the two focus knobs. It is the commonly used focus in viewing the slides.
Image 14: The stage height adjustment is at the bottom left.
- Stage height adjustment – It allows you to easily adjust the placement of mechanical stage in both horizontal and vertical path. Adjusting the knob is important as it prevents the possibility of contact between the objective lens and slide containing the specimen.
Image 15: The illumination is at the center of the base.
- Illumination – It is the light used to illuminate the slide that contains the specimen. The light comes from the base of the microscope.
Image 16: Mirror sets on top of the base.
Image 17: The field diaphragm.
- Bottom lens/field diaphragm – it is a knob used to adjust the amount of light that gets in contact with the specimen. (5, 6, 7, and 8)
How a compound microscope works/functions?
Light begins at the base of the microscope coming from the source of illumination. It travels upward through the condenser and aperture and passes through the stage. As the light passes through, the image of the specimen on the slide is picked up by the magnification of the objective lens above it. The magnification varies. After which, the light moves to the head of the microscope reaching the eyepiece and magnified by the ocular lenses.
Basically, all the parts of the microscope work together to magnify the specimen and have a clearer view. As someone who is using the microscope, it is important to learn how to properly use and adjust the microscope.
Aside from the proper use of a microscope, it is also important to keep the microscope in perfect shape and one way of doing so is by keeping it clean. (2, 5, 8, and 9)
Why is compound microscope image inverted?
A compound microscope captures an inverted image of the specimen because every time the light passes through the lens, the image’s direction is flipped. The image always ends up inverted from the original. So, if you move the sample to the left, it moves in the right direction.
Image 18: A comparison image between a simple and compound microscope.
What is the difference between a compound microscope and a simple microscope?
- Simple microscope – It is a convex lens of small focal length and its primary use is to see a magnified image of small objects.
- Compound microscope – It is an optical instrument consists of two convex lenses of short focal lengths primarily used for observing a highly magnified image of minute objects.
- Simple microscope – It has a convex lens. It uses only one lens to magnify objects. An example of a simple microscope is a magnifying glass.
- Compound microscope – It has two convex lenses. It is called a compound microscope because it compounds the light as it passes through the lenses to magnify. The image of the object being viewed is enlarged because of the lens near the object. An eyepiece, an additional lens, is where real magnification takes place. The lens of the eye piece magnified the already enlarged image making it larger and clearer. (2, 4, and 6)
The focal length is the distance between the lens and its focus.
- Simple microscope – A simple microscope has a short focal length.
- Compound microscope – They eyepiece makes the focal length longer and more precise. The objective lens and the eyepiece make the object larger and more defined.
- Simple microscope – It has a maximum magnifying power of 10. As with the nature of magnification, a simple microscope has a fixed magnification. It magnifies the image to a certain degree that the lens allows.
- Compound microscope – It has the maximum magnifying power of 1000. A compound microscope’s magnification can be multiplied because it has an additional lens. You can magnify to the lens the highest capacity making the image clearer and more defined. (7, 9, and 10)
Presence of condenser lens
- Simple microscope – Absent
- Compound microscope – Present
- Simple microscope – Natural
- Compound microscope – Illuminator
- Simple microscope – Concave reflecting
- Compound microscope – It has both plain and concave type mirror.
- Simple microscope – For simple/basic use.
- Compound microscope – A compound microscope is commonly used for professional research purpose.
Check the table below for a detailed comparison between a simple microscope and a compound microscope.
Point of comparison Simple microscope Compound microscope Definition A convex lens of small focal length and its primary use is to see the magnified image of small objects. An optical instrument consists of two convex lenses of short focal lengths primarily used for observing a highly magnified image of minute objects. Lenses A convex lens with only one lens to magnify the object. Two convex lenses Focal length short focal length Longer and more precise Magnification Fixed magnification Varies Presence of condenser lens Absent Present Source of light Natural Illuminator Type of mirror Concave reflecting Both plain and concave Magnification adjustment No Yes Usage/application Simple/basic use Professional research purpose
Compound microscope types
Compound microscopes are categorized into four types. They are the following:
Image 19: A toy compound microscope.
- Toy microscopes – These are compound microscopes sold in toy shops. You can easily spot a toy microscope because it comes with many accessories which include but not limited to the following:
- Plastic pipettes
- Prepared specimens
- Ridiculously high magnification
A toy microscope does not have objectives manufactured as per the 160 mm standard. It has a low resolution and it is extremely difficult to achieve focus because its parts are made of plastics.
Another distinct characteristic of the toy compound microscope is its low field of view and low brightness.
Image 20: A compound microscope typically used in schools.
Picture Source: medpro-microscope.com
- This type of compound microscope is small, which makes it a portable device. So, students can bring it with them anytime and anywhere. Its eyepiece has 10x magnification and comes with three objectives: 4x, 10x, and 40x.
- Its light source comes from halogen or LED. Some even have a battery, which enables you to use the microscope even with no power supply. This is the best microscope for an amateur user.
- It is easy to use and adheres with the DIN standardized objectives. Its body is usually made from metal. Some educational/student compound microscopes have a condenser with a diaphragm for you to easily control the resolution, contrast, and depth of field.
Image 21: A compound microscope typically used in the laboratory setting.
Picture Source: ssl-images-amazon.com
- It is bigger in size when compared with the student microscope. It is also heavier. It is primarily used for laboratory setting. It comes with a wide body and base.
- Its distinct parts include a condenser, illumination, focus lock, mechanical stage, and a revolving nosepiece which can hold up to five objectives. It usually has a binocular head, which makes long-term observation easy.
Image 22:An example of a research compound microscope.
What is a Light Microscope? (with pictures)
A light microscope, also called an optical microscope, is an instrument to observe small objects using visible light and lenses. It is a highly used and well-recognized microscope in the scientific community. The device can be used to view living or dead samples and can maximize these samples up to one thousand times (1,000x) their actual size. Light microscopes include almost all compound and stereo microscopes.
This type of microscope is composed of an objective lens, an ocular lens, a stage, a light source, a condenser, a tube, an arm to support the tube, and a focusing system. The specimen is set on the stage, a platform usually equipped with metal arms to hold the specimen or slide in place. The light bulb is situated beneath the stage so that the light shines up through the specimen. The tube focuses down on the stage so that the ocular lens, or eyepiece, is at the far end of the tube and the objective lens is at the end closer to the specimen.
The objective lens is a small, round piece of glass that collects the light from a small area of the specimen at a short focal length and directs the light into the tube. The image is then magnified by the ocular lens, which is put up to the eye. Because the objective lens is convex, it focuses and directs light into its center. By contrast, the concave shape of the ocular lens serves to spread out the light as it meets the eye, thereby making the image bigger. The condenser is a lens, often implanted into the stage or located just below it, that condenses the light rays from the light source onto the spot that is being examined on the specimen above.
A simple light microscope uses only one magnifying lens, but today, most microscopes use two or more lenses to magnify the image. Most microscopes today are compound microscopes that use more than one magnifying lens. The eyepiece typically magnifies to 2x, 4x, or 10x actual size and the ocular lens may magnify 4x, 5x, 10x, 20x, 40x, 50x and 100x. A microscope usually comes with three ocular lenses of different magnification levels set on a rotating nosepiece. There may also be a fourth lens used for oil immersion viewing of specimens, wherein a drop of oil is set on the slide to further refract light and the oil immersion lens is lowered until it touches the oil droplet.
The relationship of glass to magnification and the concept of lenses were discovered by the Romans in the first century, A.D. Lenses were eventually put to use at the end of the 1200s as spectacles. This may have set the stage for Zaccharias and Hans Jannsen, Dutch spectacle makers who, in the year 1590, are said to have invented the first compound microscope by experimenting with several lenses in a tube. The validity of the Jannsens’ claim to this invention, however, is highly disputed. Many historians credit Tuscan scientist Galileo Galilei with the development of the compound microscope and technologically similar telescopes in the early 1600s.
Later, a Dutch store apprentice named Anton Von Leeuwenhoek refined lens making to achieve a steep curvature on a small lens, allowing him to focus on much smaller specimens than ever before. He is often referred to as a father of microscopy as he introduced the microscope as a vital instrument to the field of biology. In addition to other discoveries, Anton Von Leeuwenhoek was the first to view bacteria, yeast, and the organisms in a drop of water.
Can one use a hemocytometer on a compound upright microscope? - Biology
Stephen M. Wolniak
Department of Cell Biology & Molecular Genetics
University of Maryland
College Park, Maryland, 20742
Teaching Interests - Microscopy
I provide the information presented below for students who generally know little about the basics of image formation in the light microscope. If you wish to use this information, kindly credit (blame) me for the effort it took to generate the document.
Principles of Microscopy
The microscope that is available to you for general use in this laboratory is a sophisticated optical instrument that can provide you with high-resolution images of a variety of specimens. Image quality is based largely on your ability to use the microscope properly. Below you will find some basic information that you have probably heard before, but information that is rarely presented in a thorough way.
The magnification of small things is a necessary facet of biological research, but the fine detail in cells and in subcellular components requires that any imaging system be capable of providing spatial information across small distances. Resolution is defined as the ability to distinguish two very small and closely-spaced objects as separate entities. Resolution is best when the distance separating the two tiny objects is small. Resolution is determined by certain physical parameters that include the wavelength of light, and the light-gathering power of the objective and condenser lenses. A simple mathematical equation defines the smallest distance (dmin) separating the two very small objects:
dmin = 1.22 x wavelength / N.A. objective + N.A. condenser
This is the theoretical resolving power of a light microscope. In practice, specimen quality usually limits dmin to something greater than its theoretical lower limit.
N.A. (Numerical Aperture) is a mathematical calculation of the light-gathering capabilities of a lens. The N.A. of each objective lens is inscribed in the metal tube, and ranges from 0.25-1.4. The higher the N.A., the better the light-gathering properties of the lens, and the better the resolution. Higher N.A. values also mean shorter working distances (you have to get the lens closer to the object). N.A. values above 1.0 also indicate that the lens is used with some immersion fluid, such as immersion oil.
From the equation above, you should be aware that the N.A. of the condenser is as important as the N.A. of the objective lens in determining resolution. It is for this reason that closure of the condenser diaphragm results in a loss of resolution. In practice, at full aperture and with good oil immersion lenses (N.A. 1.4 for both the condenser and the objective) it is possible to be able to resolve slightly better than 0.2 µm. From the equation above, it should also be clear that shorter wavelength light (bluer light) will provide you with better resolution (smaller dmin values). However, there are practical considerations in how short the wavelength can be. In the early 1950's, a UV microscope was designed, but required quartz objectives and a specialized imaging device. The quartz lenses provided slightly better resolution (dmin = 0.1 µm), but image quality suffered from an inability on the part of the manufacturers to correct for aberrations caused by the quartz. The human eye is best adapted for green light and our ability to see detail may be compromised somewhat with the use of blue or violet. Most manufacturers of microscopes correct their simplest lenses (achromats) for green light.
- Magnification and Imaging -
Most microscopes in current use are known as compound microscopes, where a magnified image of an object is produced by the objective lens, and this image is magnified by a second lens system (the ocular or eyepiece) for viewing. Thus, final magnification of the microscope is dependent on the magnifying power of the objective times the magnifying power of the ocular. Objective magnification powers range from 4X to 100X. Lower magnification is impractical on a compound microscope stand because of spatial constraints with image correction and illumination. Higher magnification is impractical because of limitations in light gathering ability and shortness of working distances required for very strong lenses. Ocular magnification ranges are typically 8X-12X though 10X oculars are most common. As a result, a standard microscope will provide you with a final magnification range of
Each objective lens consists of six or more pieces of glass that combine to produce a clear image of an object. The six or more lenses in the objective lens are needed to provide corrections that produce image clarity. The interaction of light with the glass in a lens produce aberrations that result in a loss in image quality because light waves will be bent, or refracted, differently in different portions of a lens, and different colors of light will be refracted to different extents by the glass. Spatial aberrations (e.g., spherical aberration) can be corrected by using lenses with different curvature on their surfaces, and chromatic (i.e., color) aberrations can be minimized by using multiple kinds of glass in combination. These corrections increase the cost of the lens to the extent that an apochromatic objective lens exhibiting full color correction and extremely high N.A. can cost several thousand dollars. This objective lens is about the size of your thumb.
The objective lenses in most microscopes are achromats, and best suited for imaging with green light. Green filters narrow the bandwidth of the light, and make achromat objectives reasonably effective for most routine uses. The achromat lenses are not suitable for critical high-resolution imaging with white light, because red and blue light do not focus in the same plane as green light. Chromatic aberrations will degrade resolution in color images obtained with achromatic objectives. Color photomicrography aimed at the highest level of resolution and image clarity should be performed with totally corrected apochromatic objective lenses. Fluorite lenses, offer intermediate levels of correction, better than achromats but not as good as apochromats. Fluorite lenses are well suited for fluorescence microscopy because of their high transmittance of shorter wavelength light. Higher levels of correction make objective lenses more expensive the price range for apochromatic objectives goes from about $3,000 to over $10,000.
The oculars in most microscopes are designed to work optimally with the objective lenses from the same manufacturer. Each manufacturer makes some of the color and spatial corrections in the objective and the remainder of the corrections in the ocular. Mixing brands will usually result in a degraded image. In addition, when you look into a microscope, the magnified and corrected image you see through the oculars is actually a virtual image (as opposed to a real image). The ocular, designed to provide a corrected virtual image when viewed by eye, is not suitable for the generation of photographic or video images through the microscope. For photography or video microscopy it is necessary to use a projection lens that generates a corrected real image. Many of the newer microscopes provide total image corrections in the objective lens, thus obviating many of the concerns aboout matching glass components from the same manufacturer. Nevertheless, it is a good practice not to mix parts from one manufacturer with those of another, because unintended image degradation can result.
An essential factor in producing a good image with the light microscope is obtaining adequate levels of light in the specimen, or object plane. It is not only necessary to obtain bright light around the object, but for optimal imaging, the light should be uniform across the field of view. The best way to illuminate the specimen involves the use of yet another lens system, known as a condenser. The front element of the condenser is usually a large, flattened lens that sits directly beneath the specimen. Its placement on a movable rack provides you with the means to focus the light beam coming past the object and maximixe the intensity and control the uniformity of illumination. Two apertures in the illumination system allow you to regulate the diameter of the illumination beam by closing or opening iris diaphragms. One of these diaphragms, housed within the brightfield condenser and known as the condenser diaphragm, allows you to increase contrast, but at the cost of worsening resolution. The second of these diaphragms, known as the field aperture diaphragm, does not affect resolution as dramatically and is regularly adjusted for optimal illumination.
Optimal illumination of a specimen with all microscopes currently manufactured is achieved by using a variation of Kohler Illumination, where (for those of you are technophiles) the filament of the light source is in focus at the rear focal plane of the objective lens. Operationally, it is easy to obtain optimal illumination for brightfield (or phase contrast) by first placing any specimen on the stage and focusing on the object. Next, turn the ring for the field aperture diaphragm (the lowest aperture on the microscope) so that its edges obscure the periphery of the field of view. Next, raise or lower the condenser until the edges of the field aperture diaphragm are clearly focused. Do not refocus the objective on the specimen while you are adjusting the condenser. It may be necessary to center the field aperture diaphragm, using the condenser centering screws. When the microscope is properly illuminated, both the object and the edges of the field aperture diaphragm should be in the same plane of focus and the field iris diaphragm should be centered in the field of view.
Phase Contrast Microscopy
The human eye can perceive changes in light amplitude (intensity). Unstained biological specimens, such as living cells, are essentially transparent to our eyes, but they interact with light in a fairly uniform way, by retarding (slowing) the passage of a light beam by approximately 1/4 of a wavelength ( />). By slowing a light beam this much relative to another light beam that had passed though the surrounding medium, the biological specimen alters the phase of the beams. Intensity (amplitude) is additive and light rays that are 1/2 />out of phase are perceived as darkness. Zernicke realized that if he could retard the light passing through biological specimens without affecting the light passing through the surrounding medium, he could generate changes in amplitude within living cells. The phase contrast microscope was invented by Zernicke in the 1930's as a means to generate contrast in biological specimens, changing these invisible phase differences into visible amplitude differences.
Zernicke employed an optical trick to separate the light beams interacting with the specimen from those that do not encounter the specimen. To separate the beams of light from each other, he placed a transparent ring (known as an annulus) in an opaque disk and inserted this disk into the optical path of the microscope, within the condenser. He placed a complementary ring inside the objective lens. Nearly all of the light that passes through the sample but misses the specimen then passes through the objective lens through this ring. Most of the light that passes through the specimen is scattered and some of it enters the objective lens in such a way that it will not pass through the objective lens ring, but will pass this plane in some other location. He designed the glass plate holding the ring so that all light missing the ring would encounter an additional 1/4 />of retardation relative to the beams of light that had not interacted with the specimen, placing the light rays that had interacted with the specimen out of phase with rays that had not interacted with the specimen by 1/2 />. He found that a reduction in intensity of the light that had not passed through the specimen would create a grey background and increase contrast even more, with some parts of the specimen darker and other parts of the specimen brighter than the background.
The operation of any microscope in the phase contrast mode requires that you first set up proper brightfield illumination, with a centered field iris diaphragm whose edges are in focus in the specimen plane. Next, rotate the condenser turret cylinder until the number on the condenser turret matches the number engraved on the objective lens. Under this condition, the condenser annulus is matched to the phase ring present in the objective. Next, remove one of the oculars and insert the Bertrand focusing telescope into the ocular hole. This lens enables you to see the rear focal plane of the objective lens, the plane where the ring resides. You will see a bright circle of light (the condenser annulus) and a dark ring (present within the objective). The dark ring is stationary, but the bright annulus is not. You may need to align the annulus with the ring so that the two are superimposed. On the back side of your condenser, you will find two adjustment screws that permit this alignment to be performed. When the ring and the annulus are aligned, place the ocular back into the microscope. The difference between phase contrast and brightfield for the observation of living cells is significant.
In certain classes of atoms and molecules, electrons absorb light, become energized, and then rapidly lose this energy in the form of heat and light emission. If the electron keeps its spin, the electron is said to enter a singlet state, and the kind of light that is emitted as the electron returns to ground state is called fluorescence. If the electron changes its spin when excited, it enters the triplet state, and the kind of light that is emitted as the electron returns to ground state is known as phosphorescence. Phosphorescence is much longer-lived than fluorescence. Both fluorescence and phosphorescence emissions are of particular wavelengths for specific excited electrons. Both types of emission are dependent on specific wavelengths of excitation light, and for both types of emission, the energy of excitation is greater than the energy of emission. Described another way, />of excitation light is shorter than />of emission light. In biology, we can utilize fluorescence in localization reactions, to identify particular molecules in complex mixtures or in cells. Fluorescence has the advantage of providing a very high signal-to-noise ratio, which enables us to distinguish spatial distributions of rare molecules. To utilize fluorescence, we need to label the specimen (a cell, a tissue, or a gel) with a suitable molecule (a fluorochrome) whose distribution will become evident after illumination. The fluorescence microscope is ideally suited for the detection of particular fluorochromes in cells and tissues.
The fluorescence microscope that is in wide use today follows the basic "incident-light" design of Ploem, who employed a novel arrangement of filters with a chromatic beam splitter (often wrongly called a dichroic filter both by biologists and microscope sales people). With the incident light fluorescence microscope, the object is illuminated with fluorescence excitation light through the objective lens. The object emits longer-l fluorescence in response to the shorter- excitation light. The objective lens then serves both for illumination and imaging. The chromatic beam splitter transmits or reflects light, depending on its color. For this application, shorter light is reflected and longer light is transmitted by the splitter. Ploem placed the chromatic splitter in the optical path between the objective lens and the ocular, at a 45° angle, so that it would reflect shorter light downward toward the objective. The longer- fluorescence emission light would be transmitted through the chromatic beam splitter toward the ocular.
The microscopes that you have utilized in this and other courses all operate in the same general fashion. Light beams pass through a condenser lens system and provide illumination of an object at many points simultaneously. For incident light fluorescence microscopy, the objective lens also acts as a condenser for the excitation light beam. In its interaction with the object, some of this light is absorbed, some of this light is scattered, some of this light is reflected, and some of this light is slowed or retarded (relative to a beam of light that does not pass through the object). A portion of the light that has interacted with the object then passes through the imaging lens system of the microscope where it provides us with visual or pictorial image information about the object. Like the process of illumination, the process of image generation operates in a parallel fashion, where large numbers of light beams contribute to the image simultaneously. Resolution is limited by the closeness of overlapping points of brightness or darkness. In a practical sense, the limit of resolution is 0.18-0.2 µm with the best available objective lenses and a good specimen.
To observe cells with the fluorescence microscope, it is important to know the spectral characteristics of the fluorochrome that has been employed. In order to excite the fluorochrome properly and then observe its fluorescence emission, the appropriate filter packages must be present in the microscope. The fluorochrome may not fluoresce at all if the cells are illuminated with the inappropriate filter pack present in the optical path. Finally, for any kind of fluorescence localizations to be performed, it is essential to have the appropriate controls, to be sure that the cells do not exhibit excessive autofluorescence (that is, they do not glow in the absence of the fluorochrome), and that the fluorochrome is responsible for the localization pattern observed. In the laboratory, we have several microscopes equipped for incident light fluorescence microscopy.
Confocal Scanning Optical Microscopy
In the incident light fluorescence microscope, a light beam passes through a chromatic beam splitter and then the objective lens to illuminate a specimen. This light beam is used to excite electrons in fluorochrome molecules present in the object. As some of those excited electrons return to their ground state, the emission of light is detectable through the oculars of the microscope, or with a camera or video printer. The image is generated continuously, across the entire field of view. A primary problem with the fluorescence images generated in this way is that out-of-focus fluorescence appears as 'flare' in the object, and reduces the signal substantially. In addition, human eyes are not sufficiently sensitive photodetectors for the lowest levels of fluorescence, and most video-based imaging systems are only slightly better than your eyes. Under conditions where there is sufficient signal for you to easily observe fluorochrome distribution patterns, the excitation light can be of sufficient intensity to photooxidize (i.e., burn) your specimen. Much information can be lost with just a few seconds of exposure to the excitation lamp. The Confocal Scanning Optical Microscope, an expensive piece of instrumentation that illuminates the object with a small beam of light in a point-by-point (i.e., serial) fashion, eliminates most of the photoxidation problems, permitting the observation of objects for extended periods at very high resolution with little loss of signal. The placement of a small aperture in the beam path generates a small depth of field, and effectively eliminates out of focus information in image formation.
The confocal scanning optical microscope is designed to illuminate an object in a serial fashion, point by point, where a small beam of light (from a LASER) is scanned across the object rapidly in an X-Y raster pattern. The raster pattern can be created in several ways, but in one of the more popular instruments, it occurs as a consequence of the simultaneous rotation and vibration of a polygonal mirror. The vibration is caused by the activity of a servogalvanometer, while the rotation is caused by the activity of a small electric motor. Thus, a bright spot of light scans across an object from top to bottom, line by line. The image is also generated point-by-point. Image formation is translated into intensities at each spot in the X-Y raster by a photomultiplier tube. The intensity information is digitized and stored in a computer. A complex image processing software package permits visualization and manipulation of the images. Resolution is limited by spot size for the LASER and approaches 0.12-0.15 µm for an ideal specimen and with the best available objective lenses.
The manufacturers of confocal scanning optical microscopes include a pinhole diaphragm at a very special place in the optical path, near to the site of the photomultiplier tube. This pinhole is situated in a plane where the light from the in-focus part of the image converges to a point. Light from object planes above or below that of the focused image do not converge at the spot in the optical path occupied by the pinhole. Because of this design, out of focus image information is darkened to the extent that it is not detectable. The consequence is that all out of focus information is removed from the image and the confocal image is basically an 'optical section' of what could be a relatively thick object. The 'thickness' of the optical section may approach the limit of resolution, but in practice, the resolution in the Z-direction is somewhat greater, approximately 0.4-0.8 µm. The value of optical sectioning is best realized with fluorescence microscopy, where out-of-focus information alters, distorts, or even degrades the image. Because the confocal images are stored in a computer, it is possible to stack them up and generate three-dimensional reconstructions. The image processing programs also enable us to rotate these images and observe three-dimensional aspects of cellular structure. It may be clear to you that the computer responsible for these image manipulations must be fast and powerful. The biggest problem is one of image storage, where single images can routinely occupy >1,000,000 bytes of space. In rather short periods of use, it is easy to accumulate sufficient numbers of images to fill the largest of hard disks.
Two of the three the confocal scanning optical microscopes located on campus were manufactured by Carl Zeiss, located in Germany. The newest instrument (model 510) has three lasers and four photomultipliers and is designed so that we could illuminate with two or three colors of light in rapid succession and detect as many as three superimposed signals (essentially) simultaneously. The signals are separated from each other on the basis of color, using an acoustical optical tunable filter (AOTF). The optical microscope is an inverted stand. The most important operational difference between this microscope and the upright microscope in most laboratories is that with this instrument, the slide is placed in the stage holder upside-down. Like most modern research microscopes, this microscope is equipped for phase contrast, differential interference contrast and fluorescence microscopy and can be used with these imaging techniques for conventional imaging. However, it is equipped with a number of very highly corrected (read expensive) objective lenses attached to the turret, just below the stage. These lenses are necessary for high resolution confocal microscopy. The confocal part of this microscope is contained in a box that is attached to the inverted stand through an access port. As is the case with incident light fluorescence, the laser light passes through the objective lens to illuminate the specimen. An air suspension table is designed to eliminate vibrations present in the building.
Deconvolution Microscopy and Image Reconstruction
An alternative approach for eliminating flare from fluorescent image stacks is to perform intensive, iterative image analysis and processing, from objects that have been illuminated and photographed at multiple, adjacent focal planes. The images are obtained with a high-performance CCD camera, operating at very high magnification, using standard incident light fluorescence microscopy. The excitation source is a mercury arc lamp, and bandwidth for excitation and emission are controlled by filters placed in rotating filter wheels. The lamp is stabilized and the beam is randomized for uniform illumination of the specimen. Unlike confocal scanning instruments, the whole field of view is illuminated simultaneously with this microscope. It is possible to perform rapid sequential imaging (4 colors) from multiple fluorochromes with this microscope. At very high magnification, fluorescence from any spot in a cell acts as a point source. By knowing the image spread functions above and below the plane of focus, it is possible to determine points of origin for fluorescence, and spreading beams of light from that point source, above and below the plane of focus. An iterative algorithm, which is essentially a linear combination is performed by a computer on the adjacent pixels within a single image plane, and in successive image planes through the thickness of the object. Spreading light beams are subtracted from reconstructed image stack, and that light is added back to the source, thereby reducing noise and increasing signal, respectively. We have recently acquired a sophisticated DeltaVision microscope from Applied Precision, Inc., which is designed to acquire these images and then perform the computer-intensive operations. This kind of microscope is particularly well suited for generating three-dimensional fluorescence images from small, living cells.
Polarization Light Microscopy
When light passes through an object, it interacts with some or all of the atoms and molecules present in that object. In these interactions, sometimes light of a particular (i.e., color) is absorbed by the atoms or molecules, while sometimes light is scattered. The interaction of light with a translucent object often results in a slight reduction in the velocity of the light beam. The extent of this reduction in velocity can be measured as the refractive index of the object. For certain kinds of objects, especially those with high order in particular axes of the object, such a crystalline or paracrystalline arrays, the interaction with light beams is vastly different, depending on the orientation of the object relative to the impinging light beam. As a result, the refractive indices are measurably different in different axes of the object. Such an object with multiple refractive indices is termed birefringent. Birefringence (multiple refractive indices) results from the alignment of atoms or molecules in particular planes of an object these atoms or molecules interact strongly with light beams impinging on them from a particular direction, and to a far lesser extent with light beams impinging on them from a different direction. There are two kinds of birefringence, intrinsic birefringence, which results from atomic or molecular order in a crystalline or paracrystalline array (i.e., calcite crystals, membranes) and form birefringence, which results from supramolecular associations in paracrystalline arrays (i.e., microtubules in a spindle).
- Polarized Light and Birefringent Retardation -
Any light beam shining in a particular direction vibrates in all directions around the axis of travel. Light beams whose vibration has been restricted to a single plane, or to a few planes is known as polarized light. Birefringence is directly observable as differences in intensity in different axes of crystalline or paracrystalline objects when they are viewed with polarized light. Since birefringence results from differences in the number of interactions between the light beam and atoms or molecules in the object in different directions, in practice, the object is rotated around the plane of vibration for the polarized light beam to maximize the intensity differences in the object (usually, the dominant object axis is at a 45o angle relative to the plane of polarization). The extent of the difference in refractive indices in different axes of the object is a measurable quantity known as birefringent retardation (BR). BR is measured (as a distance) by placing an object with known birefringent retardation into the light beam, and, by rotating the calibrated object around the optic axis, extinguishing the brightness in the sample. Using this compensation technique, BR has been shown to be directly related to the number of aligned microtubules in mitotic spindles in living cells. This principle and procedure can be of importance in studying microtubule dynamics, where mitotic spindles of developing sea urchins can be visualized in a totally noninvasive way.
USB computer microscopes, also called computer or computer-connected microscopes, plug into a USB port on a computer or television. Instead of looking using an eyepiece, the viewer examines the specimen via the computer monitor or TV screen, like a webcam with a lens. Most of these microscopes are handheld and can save images as files or videos. However, most have only low-level magnification, and adequate illumination can be a problem.
Pocket microscopes are handheld, durable, and useful for field work. Sizes vary, and some are the size of an ink pen. Most use natural light or are battery-powered, with 25x to 100x magnification. Portable microscopes may also be digital.