Digital Microscope Comparison

There are many digital microscopes on the market, and sometimes it can be difficult to determine which microscope best fits your needs. In this post we will compare the Motic BA310 digital microscope with the Swift M10-LB digital microscope.

 

Motic Digital BA310 Microscope

Each of these digital microscopes plug into the USB port, include software and will allow you to view a live image on your computer. The Swift M10-LB offers higher mega pixels (5) if using the SD removable card however, much lower mega pixels (0.3) when viewing an image on the computer or capturing the image with the software. The Motic BA310 digital microscope offers 3 mega pixels, but does not include an LCD screen or removable SD card.

Swift M10-LB Digital Microscope

Each microscope has the option for phase contrast, although the Motic BA310 Digital microscope offers more flexibility in that there are options for a phase slider (simple phase) as well as darkfield.

Human blood cells, 400x, captured with Motic BA310 Digital microscope.

Human blood cells, 400x, captured with Swift M10-LB digital microscope.

If you have any questions regarding either of these microscopes please email us.

Walton & Beckett Reticle

Walton & Beckett reticles are used primarily for counting particles, and in particular - asbestos dust particles or glass fibers.

Generally the fibers being counted are smaller than 5 microns in size. The circle on the reticle is divided into four by two diametrical lines scaled in units of 5 and 3 microns respectively. These are the critical measurements of fiber lengths and diameters used in asbestos fiber counting. The Walton and Beckett reticle has a series of shapes that have been designed to compare with fibers.

Protein Crystallography Microscopes

A good stereo microscope is required during evaluation of crystallization experiments in order to distinguish between amorphous and crystalline specimens. A typical microscope setup would consist of a zoom stereo microscope with a cross polarizing attachment and a minimum of 10x magnification. Higher magnifications are ideal for easy detection of small microcrystals.

One of several protein crystallography microscope packages sold by Microscope World.

Protein crystallography microscopes are used to look at properties of crystals such as birefringenece (or double refraction) and for identifying conditions that cause precipitation or crystal growth. When identifying crystal growth, a polarizer would not be used because the color of the precipitate is a crucial indicator.

The microscope shown above uses pseudo-darkfield illumination and a dual-arm adjustable self supporting fiber optic illuminator. This allows top illumination from various angles and provides better light for examining crystals.

Using Polarizing Filters to Reduce Glare

When using stereo microscopes with highly reflective items, it can sometimes be helpful to also utilize a polarizing filter. The images shown below were both captured with the SMZ-168 stereo zoom microscope and a Jenoptik Speed XT Core 3 camera.

Printed circuit board captured with brightfield only.

Printed circuit board captured using polarizing filter.

Polarizing filters can especially help to reduce glare when photos are being captured with the microscope. If you do not achieve the exact image you are looking for, try rotating the polarizing filter slowly while looking through the microscope.

Microscope Immersion Oil Use

Many microscope manufacturers include a small amount of immersion oil to use with oil immersion objectives when microscopes are sold.  Since it only takes a drop to use this material, a small bottle will last for quite a while, but eventually needs replacement. 

Lenses designed to be used with immersion oil usually have a black ring around the barrel next to the colored ring identifying the magnification. They will also show “oil” in the identifying markings. These are the only lenses which should be used with oil and they should always be used with oil.  The image without the oil interface will show optical aberrations and result in a very poor image.

Air has a refractive index of 1.0 and a good oil lens has a numerical aperture of 1.25 or 1.30.  The glass used in slides and coverslips typically has a refractive index of 1.515  The full resolution of the lens can only be achieved by matching the refractive indexes of the materials the light passes through and it will be limited by the lowest number. Using air instead of oil limits the resolution to a lower numerical aperture, less than 1.0, resulting in a lower resolution and less brightness.

The point of an immersion oil objective is a superior image in terms of resolution and brightness, therefore matching the refractive indexes, 1.515 in the glass and the typical immersion oil with a refractive index of 1.515 results in the best possible image.  In this case the resolution is improved by approximately 50% over a dry lens with the same magnification.

After use the objective does not have to be cleaned completely if it will be used again within days. However,  it is a good idea to blot the front of the lens with lens tissue to remove excess oil.  If the microscope will not be used again soon, the oil should be cleaned off.  This assumes you will be using the same immersion oil.  Anytime you change brands of immersion oil, the lens should be cleaned of any of the previous oil.  Mixing oils can result in clouding of the material and a diminished image, not to mention, a new compound with unknown properties.

The biggest mistake made with the use of immersion oil is going back to a lower power objective once oil has been added to the slide.  It is a very good habit to examine your sample completely at low power before you oil the slide.  If you oil the slide for the 100x microscope objective and go back to the 40x, you will get oil on the 40x and the image will not be sharp.  It is much easier to keep oil off the 40x than it is to do a good cleaning after it gets oiled.

Letting oil remain on an objective not designed for oil also increases the potential for the oil to degrade the seal or the lens cement and seep into the interior of the lens assembly.  This very effectively ruins the objective which in most cases has to be replaced.

Fluorescence and Autofluorescence

Epi fluorescence is one of the most common techniques used in scientific research today. Samples are "tagged" with certain compounds that attach to a specific part of the tissue or cell and then exposed to a discrete wavelength of light to "excite" the compounds. The wavelength of light used is matched to specific compounds, or fluorochromes, so that the resulting image only shows the structure that has been labeled. For example, a very common fluorochrome is DAPI, which is used to attach to the nucleus of a cell. The wavelength of light used to excite DAPI is 325-375 nanometers (nm), which is in the ultraviolet range. When the DAPI is excited by this wavelength of light, it releases energy at a longer wavelength, in this case 435-485nm, which is blue light. So the sample is illuminated in the ultraviolet, but the color that is reflected back to the eyepieces is in the blue. When you see images of cells in literature, they are almost always blue.

Other fluorochromes all have a distinctive color associated with them. FITC is excited by blue light, but reflects back green so FITC images will be green. TRITC is excited by green light, but reflects back orange-red, so those images will be in the orange-red range.

Some material, especially plant material has the inherent ability to fluoresce, or react to specific wavelengths of light, without having any dyes or fluorochromes attached to the material. This is called "autofluorescence" and can interfere with using this research technique effectively. It is however, helpful for other tasks, such as using a plant sample when you are working to align the fluorescence illumination. This autofluorescence does not fade or quench as do biological samples so you are not damaging your cells while performing the alignment. It also allows practice in setting up photography without damaging the sample. And the bonus is that it results in beautiful images!

Porton Counting Reticles

A porton counting reticle is used to count and measure abrasives and dust particles - most commonly asbestos, paint particles or varnish particles.

In order to determine if dust particles in the air are at a safe level, a sample is collected on a filter that will reflect the concentrations of dust at the time and place of sampling. Once the sample is collected, it is placed under a phase contrast microscope and examined using a porton counting reticle. This must be done within two days of sample collection.

The half of the Porton reticle with vertical lines serves to define the counting area of the field. Twenty fields located at random on the sample are counted and total fibers longer than 5um are recorded. Any particle having an aspect ratio of three or greater is considered a fiber. The fiber lengths are determined by comparison with the cirles on the reticle. The square that is divided into six rectangles is the counting field area.

You can learn more about Porton Counting Reticles here.

Microscope Condenser and Aperture Diaphragm

When first looking at a microscope, it can sometimes be confusing if parts and controls are located in different places. However, there are always certain components that are present on a microscope. Eyepieces, objectives, and focusing controls are easy to find, but aperture and field diaphragms may not be. Field diaphragms are useful to adjust for correct microscope alignment, but are not always included in less expensive microscope models.

The aperture diaphragm (also called an iris diaphragm) controls contrast, and is found in the condenser, which sits right below the stage in line with the microscope objectives. The condenser may be movable, both in the horizontal and vertical directions. If the condenser is fixed and has no position adjustment, it has been pre-centered at the factory, but it should still have an aperture diaphragm with a movable collar or knob. There may be markings on the body of the condenser to correspond with the varying numerical apertures of the objectives used but with a bit of practice, you can make adjustments by viewing the image quality.

The collar or slider adjusts the degree the aperture is open or closed, thereby affecting the depth of field, usable numerical aperture and overall image quality. The goal of correct adjustment is a balance between the best resolution and good image contrast. If the aperture is wide open the image will appear washed out with no contrast and detail will be difficult to observe. If the diaphragm is closed down too much, the image will be "grainy" with much less resolution and addition of "artifacts" (dust and debris) into the image.

Image of tonsil tissue captured with the aperture diaphragm open. Captured at 40x magnification with the Swift M10 digital microscope with built-in camera.

Image of tonsil tissue captured with the aperture diaphragm correctly adjusted. Captured at 40x magnification with the Swift M10 digital microscope with built-in camera.

Image of tonsil tissue captured with the aperture diaphragm closed down. Captured at 40x magnification with the Swift M10 digital microscope with built-in camera.

The aperture should be closed down approximately 1/2 to 1/3 for proper use. You can take out one of the eyepieces and look down the eyepiece tube to see the opening and closing of the aperture diaphragm while adjusting it. The more practical method of adjustment is to open the diaphragm completely and then slowly close it down while viewing the specimen. As soon as you see the contrast improve, leave the diaphragm at that setting. It is a good idea to remove one of the eyepieces and note how much the diaphragm is closed down. You will soon recognize the best position without pulling out the eyepiece.

This microscope aperture diaphragm adjustment is critical to achieve the optimal image of the sample. A poor image due to misalignment of the microscope is very noticeable and hard to explain away. Take time to work the aperture diaphragm on your microscope to optimize your image - it is time well spent.

Phase Contrast

Most biological samples are clear or transparent and need to either be stained or viewed with contrast enhancement techniques in order to be visible under the microscope. In 1938 Frits Zernike, a Dutch mathematician and physicist, developed what is now referred to as a phase contrast microscope to allow these types of specimens to be seen and examined. He modified both the lens and the condenser by putting a ring in both and aligning these elements so that the rings lined up. This caused a phase shift of the light where it went through the sample resulting in a detailed image of the specimen.

All microscope manufacturers now product phase contrast microscopes for a variety of applications based on this design. Almost all inverted, or tissue culture microscopes, use phase contrast or other contrast enhancement techniques since samples are typically tissue samples or live cells. Live cell samples cannot be stained without killing the sample. Other techniques to do this are Differential Interference Contrast, referred to as DIC and Hoffman Modulation Contrast, both of which are considerably more expensive.

Phase contrast alignment is simple but critical, as seen in the cheek cell images shown below. The technique to accomplish this can be found here.

These are cheek cells under a brightfield microscope at 40x.

This is the exact same slide as above with phase contrast added.  This image was captured using the Motic AE31 inverted microscope with a 40x LWD phase contrast objective lens and the Lumenera DK1-CCD monochrome 1.4 mega pixel CCD microscope camera.

This is the same slide with the phase rings misaligned.

This is the same image with the wrong phase ring used.

Biological vs. Metallurgical Microscope Objectives

Compound microscopes are divided into two major categories depending on the type of sample and the light path used to examine it. Typical biological microscopes are designed to look through the sample, with the light shining up through the specimen - this is referred to as diascopic illumination. The samples are very thin and transparent or translucent.

Opaque specimens are examined with light from above the sample that is reflected back - this is referred to as episcopic illumination. These samples are viewed with metallurgical microscopes in metallurgy, engineering, manufacturing and other industrial applications.

The objectives for these two categories are distinct and are marked on the barrel of the objective. The objective shown below is for reflected light. The BD indicates that this microscope objectives is intended for brightfield / darkfield illumination, a standard technique in reflected light microscopy. (BD reflected light objectives also usually have a larger thread mount and barrel to accommodate the additional light path. These lenses will not fit the standard nosepiece). EPI indicates that this microscope objective is used for epi-illumination. Immediately after the infinity symbol there is "/0" - this means the objective is intended to be used without a cover slip. This is also the main difference in the design of reflected light microscope objectives.

Biological objectives, unless noted NCG (no cover glass) or water immersion, will be designed to include a cover slip between the specimen. After the infinity symbol the marking usually will say "/0.17" - this is the standard thickness of a cover slip. If using a fixed tube length system, the markings will show "160/0.17" or "170/0.17". If the sample has no cover slip it will not be noticeable at low power of 4x and 10x. However, at 40x and above this 0.17 is crucial and the resulting image is very poor without the cover slip in place.

A fixed tube length microscope objective.

There may be a few applications when these objectives cross the line. So keep this in mind if working with a non-traditional sample.

Viewing Wires with a Microscope

Microscope World recently had a customer who needed to view wires with a microscope, and in particular they were interested in measuring the wire casing.

Using the National Optical DC5-420TH digital stereo microscope, these images were captured and measurements made with the included microscope software.

Wires captured at 10x magnification.

20x magnification.

30x magnification.

40x magnification.

The small white text box you see in each image notes the section that was measured, and how long it is. For this project the text boxes were kept small on purpose, but it should be noted that these can be made much larger for ease of reading. Additionally, the measurement feature allows you to measure not only lines, but area, diameter, etc. You can learn more about the software here.

Fluorescence Microscopy

By using fluorescence or phosphorescence, a fluorescence microscope allows the study of organic or inorganic substances. Whereas traditional light microscopes use only reflection or absorption to view specimens, fluorescence microscopes illuminate a sample with light of a wavelength that causes fluorescence in the sample. This light is a longer wavelength than the illumination, and is then detected through the microscope objective.

Two filters are typically used in fluorescence microscopy:

1. An illumination (excitation) filter – ensures the illumination is near monochromatic and at the correct wavelength.
2. Second emission (detection) filter – ensures none of the excitation light source reaches the detector.

Fluorescence microscopy takes a different approach to creating a light microscope image to transmitted or reflected white light techniques such as phase contrast or differential interference contrast. These two contrasting optical microscopy methods provide very different, but complementary data.

So how does a fluorescence microscope work?

The specimen is illuminated with light of a specific wavelength that is absorbed by the fluorophores, causing them to emit light of longer wavelengths (in other words a different color than the absorbed light). The illumination light is separated from the weaker emitted fluorescence through the use of a spectral emission filter. Typical components of a fluorescence microscope include:

1. Light source (xenon arc lamp or mercury-vapor lamp)
2. Excitation filter
3. Dichroic mirror (or dichromatic beamsplitter)
4. Emission filter

The filters and dichroic are chosen to match the spectral excitation and emission characteristics of the fluorophore (color) used to label the specimen. The distribution of a single fluorophore is imaged at one time.

Most fluorescence microscopes are “epi-fluorescence microscopes”.  Epi refers to the fact that the excitation and observation of the fluorescence are from above the specimen.

 Fluorescence microscope image of mouse embryo.