Microscopy: advances in scientific research and education (A. Méndez-Vilas, Ed.)
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Microscope maintenance and quality control: A practical guide
Brady Eason1,2,*, David Young1,*, Aleksandrs J. Spurmanis1,3, Tse-Luen (Erika) Wee1, Daniel Kaufman4
and Claire M. Brown1,4,5
1
McGill University Advanced BioImaging Facility (ABIF),
Currently at University of Western Ontario Dental School,
3
Currently a Technical Representative, Nikon Canada Inc.,
4
Department of Physiology, 3649 Prom Sir William Osler, Bellini Building, Rm137a, McGill University, Montreal,
Quebec, Canada H3G 0B1
5
Corresponding author
*
Equal contribution by both authors
2
Since their inception in the 17th century, light microscopes have evolved from qualitative documentation tools into
sophisticated instruments capable of providing quantitative spatial and temporal information. As the light microscope has
evolved, so have the demands of researchers in addressing scientific questions. Simply observing a specimen is not
enough: there is a need for increasingly precise and accurate information to enable quantitative and reproducible
microscopy experiments. In order to perform quantitative microscopy there is a need to ensure that the microscope
performance is optimal and consistent. Here we present a concise, but detailed set of methods that are easily adaptable to a
variety of equipment designs and configurations. This flexibility is useful in circumstances where a variety of instruments
are to be monitored within tight time constraints in order to benchmark microscopes and ensure stable performance over
time.
This guide combines work from previously described tests and analytical tools, outlines published protocols and
introduces new procedures in the context of maintaining, troubleshooting and addressing instrument performance
problems [1-6] (http://zeiss-campus.magnet.fsu.edu/articles/basics/care.html). The procedures described include methods
for: 1) inspecting, cleaning and maintaining the optical and mechanical components of the microscope stand; 2)
monitoring the performance of ancillary equipment such as anti-vibration tables and live-cell incubators; 3) measuring
objective lens quality and instrument resolution using point spread function (PSF) imaging, analysis and quality metrics
[1]; 4) testing instrument alignment and field illumination uniformity [4]; 5) monitoring light source stability over the
short, moderate and long term [4]; 6) testing light source linearity and power using a power meter; 7) testing coregistration of imaging channels for accurate co-localization between the various channels for multi-color samples [4].
Image processing and analysis of the resulting data can be conducted using open-source software and plug-ins for ImageJ
that are readily and freely accessible to the scientific community [2], however MetaXpress and Microsoft Excel were used
extensively for the analysis presented here.
Quality assurance assessments are described using a systematic five-step format of: (a) experimental rationale, (b)
description of experimental methods, (c) illustration of representative results, (d) interpretation of results, and (e) potential
corrective measures. The procedures are useful for researchers new to microscopy, independent research laboratories,
centralized core facilities and seasoned microscopists. Careful implementation of the procedures will help ensure optimal
instrument performance and the generation of the highest quality scientific data.
Keywords: microscope; fluorescence; confocal; quality control; maintenance; quantitative; resolution; alignment; colocalization; signal-to-noise
1. Microscope stand maintenance
Optimal image formation relies on the proper functioning of the optical and mechanical components within the
microscope stand. The most common issue with optical surfaces along the optical light path is dirt and dust. Dirt and
dust cause artifacts within both the image plane and the out-of-focus planes. Gloves should be worn when performing
the procedures below to avoid skin oils from contaminating the optics. The use of checklists and/or log sheets is
encouraged in order to maximize the efficiency of workflow and to ensure that inspections are conducted and
documented in a consistent manner. Sample checklists and worksheets are available for download from the Advanced
BioImaging Facility webpage (www.mcgill.ca/abif - under the resources tab).
1.1 Transmitted light path
Maintenance of optical surfaces: Contaminants on brightfield condenser filters (e.g. blue correlation filters, polarization
filters), DIC filters and prisms, and glass windows on fluorescent filter cube turrets can be cleaned in several ways. The
first and least invasive step is to remove contaminants using a puff-duster (Giotto’s Industrial Inc., Rocket-air). Do not
use pressurized air canisters as they contain volatile organic aerosols that can contaminate, and even damage, sensitive
optical surfaces [7]. When contaminants are more difficult to remove, use clean room swabs (CB-FS751B www.cleanswabs.com) or swabs prepared by wrapping 100% pure cotton wool around a wooden applicator. Prepared
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cotton swabs (e.g. Q-tips®) should be avoided as they may contain synthetic fibers and other inclusions that can abrade
optical surfaces [7]. Minimal contact pressure and duration are recommended when wiping, beginning from the center
of the optical element and moving outward in a spiral motion so that particles are directed toward the edges. Ammoniafree lens cleaner diluted 1:5 in de-ionized water can be applied to surfaces contaminated by strongly adherent particles
or stains (e.g. SparkleTM, GlassPlusTM). Swabs should only be used once in order to prevent re-introducing contaminants
back onto the optical surface. A second cleaning of the objective lens with distilled water is recommended to clean off
any deposits left by the lens cleaner. Organic solvents and detergents containing ammonia (e.g. WindexTM) should be
avoided as these can degrade filter coatings.
1.2 Reflected light path
1.2.1 Fluorescent light source
Many modern light sources for fluorescence microscopy are alignment free. However, older mercury vapor arc lamps
must be properly aligned to avoid shading artifacts within fluorescence images. Alignment needs to be conducted after
each bulb change. Move to an open position in the objective lens turret and remove the dust cap. If an open position is
not available, remove one of the objective lenses. Place a white piece of paper or a business card on the microscope
stage. Attenuate the mercury arc lamp or move to a red filter cube to avoid any remote chance of eye exposure to
damaging UV light. An image of the arc and its reflection will be seen on the white paper. The image of the arc can be
aligned to the center of the objective lens opening using adjustment screws on the lamp housing. The reflected image of
the arc should be aligned in the center of the objective lens opening, but slightly shifted to the left or to the right of the
image of the arc. This alignment is done using the setscrews on the lamp housing for the reflecting mirror inside. Figure
1a shows a mercury lamp alignment tool with the image of the arc and the reflection well-focused and side-by-side.
1.2.2 ND filters
The incorporation of neutral density (ND) filters is recommended to attenuate the incident light from high intensity arc
lamps. This will reduce photo-damage to fixed samples and photo-toxicity to live samples during sample observation or
imaging [8] (http://micro.magnet.fsu.edu/primer/anatomy/cleaning.html). Most research-grade microscopes provide a
place in the fluorescence light path to insert a slider containing one or more neutral density filters (e.g. 1% and 10% ND
filters). Routine care and inspection of these filters is performed as described in Section 1.1.
1.2.3 Reflector filters and mirror maintenance
Inspect filters and mirrors within reflector cubes for contaminants. Remove any contaminants using a puff duster or by
carefully wiping the surface with a dry cotton swab. IMPORTANT: The application of water or other aqueous solutions
can irreparably degrade filter and mirror coatings. If absolutely necessary, a contact cleaner (Photonic Cleaning
Technologies, First Contact™) can be used to remove stains, fingerprints or other contaminants from these surfaces.
Figure 1b shows a reflector filter that has been damaged by exposure to the light source illumination. If there is a
significant loss of transmission efficiency, the damaged filter or mirror may need to be replaced. Rigorous maintenance
of filter cubes is less crucial when wide-field fluorescence is simply being used for sample observation such as part of a
confocal laser scanning microscope (CLSM).
1.3 Other Optical Components
1.3.1 The specimen
The specimen itself is a critical element in the optical path. Care must be taken to ensure that the specimen is devoid of
any contaminants that might interfere with proper image formation. Lens cleaner should be used to remove any media,
buffer salts or dirt from the sample coverslip. The sample holder in the stage insert should also be inspected and cleaned
routinely.
1.3.2 Objective lenses
Routine maintenance and careful inspection of objective lenses is of paramount importance. The Point Spread Function
(PSF) should be carefully measured on new lenses and routinely if lens damage is suspected (Cole et al Nat Protocols).
Lenses should regularly be removed and examined under a stereo dissection microscope to determine if there is any
damage or wear and tear. Figure 1c shows a lens seal that has been damaged. When the damage is severe enough,
immersion oil can seep in and accumulate underneath the objective casing. This accumulated oil can “weep” back out
onto the front lens making it difficult to clear during routine inspections. More importantly, the oil leak can seep into
the internal lens system and destroy the lens. Scratches on the front lens will introduce distortions and reduce the
effective resolution of the objective. Figure 1d shows an example where fragments of the compromised front lens seal
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are clearly visible when imaged with transmitted light. To minimize the introduction of dust inside the microscope
stand, objective lens dust caps should be in place in the nosepiece whenever lenses are removed.
1.3.3 Objective maintenance
The optical surface maintenance procedure above (Section 1.1) can be used to remove oil and dirt on the front lens of
the objective. Anhydrous ethanol may be used sparingly in order to clean more heavily contaminated surfaces.
IMPORTANT: the use of less polar organic solvents (e.g. acetone, ether, chloroform) is not recommended as these can
compromise anti-reflection coatings and/or lens seals [7]. Puff-dusting should be sufficient to remove particles adhered
to the back lens. Any additional intervention at the back focal lens is discouraged as this can lead to accidental damage
to the internal lens system of the objective [7]. A professional service provider should be consulted in the event that
extensive cleaning of the back lens is required.
1.3.4 Camera lens maintenance
The procedure in section 1.1 can also be used to clean camera relay lenses in wide-field microscope stands. As a result
of their close proximity to a conjugate focal plane, camera IR filters are susceptible to producing artifacts in both
transmitted and reflected light images when contaminated with dust or other particles.
1.3.5 Eyepieces
The eyepieces can be removed from the microscope tube so that both internal and external optical surfaces can be
inspected and cleaned. A puff duster can be used to clear dust out of the eyepiece tubes. The focusing mechanism (if
present) on the ocular lenses can be reset to their zero positions. Users can then re-adjunct them for their own use during
their individual sessions.
1.4 Image verification
Following maintenance, a stained specimen should be imaged with the microscope. After it is adjusted for Köhler
illumination, the quality of a brightfield image should be assessed both visually and with the camera and software. If
artifacts are visible in the image, the source of the contamination can often be identified by rotating or translating the
various elements in the transmitted light path. Additional maintenance of the affected surface may remove these
artifacts. In addition, DIC imaging or out-of-focus imaging can reveal contaminants in the light path. Seek technical
support from the instrument manufacturer if the affected surface cannot be identified or restored. Similar verification
should be done with a fluorescently labeled sample.
2. Ancillary equipment maintenance
2.1 Work area maintenance
The best way to avoid dust on optical surfaces is to eliminate it altogether. Regular furnace filters placed over air vents
and a floor based room air filter should be placed in the room to minimize dust. Routine sweeping and dusting of the
work area surrounding the microscope is also recommended. Sticky mats should be placed at the door to remove dust
from footwear as users enter the room. The computer keyboard and case openings and fans should be vacuumed often
to prevent dust build up. Wash the air table and work area with a mild detergent to remove spills or salt deposits.
2.2 Verification of anti-vibration equipment
Artifacts can arise in images when vibrations are translated to the sample. It is important to ensure that anti-vibration
equipment is functioning properly with the table level and adequate air-pressure in each table leg. It is also important to
set up microscope tables in areas that are at a distance from building intake and outtake vents, laser cooling fans,
elevator shafts and, if possible, walls backing onto fridge/freezer areas or busy hallways. Figure 1e shows one instance
where vibrations were seen within images because the laser fiber optic cable was resting on a laser-cooling fan. Figure
1f shows that when the fiber was moved away from the vibration of the fan, the artifacts within the image disappeared.
2.3 Incubator chamber maintenance
Previous reports have demonstrated the importance of maintaining appropriate temperature and CO2 conditions when
working with living cells or tissues [5, 9, 10]. Although a variety of commercially available incubation chambers
provide the researcher with the means to adjust the live specimen’s environment, few of these provide the ability to
independently measure and verify these parameters at the level of the sample. CO2 and temperature probes are also
commercially available and can be used to calibrate incubation equipment as required. A simple way to verify the
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accuracy of CO2 delivery is to equilibrate an aliquot of tissue culture medium (with standard phenol-red pH indicator)
for at least 1 hour in the incubator chamber. The medium should appear reddish-orange when exposed to 5-10% CO2.
Insufficient CO2 will cause the media to become alkaline or purple in color.
Fig. 1 Lamp alignment, objective lens inspection and artifacts of
system vibration. a) Imaging tool for mercury lamp alignment
showing optimal arc and reflection image position. b) A
fluorescence cube filter damaged by accumulated exposure to the
high intensity light source. c) Objective with torn casing. d)
Inspection of an objective with transmitted light reveals
contaminants within the internal lens system. e) CLSM image of
neurons within a 100 μm thick neuronal slice culture acquired with
a 63x/0.9 NA water immersion objective with distortions because
the laser fiber-optic was resting on a cooling fan. f) Same imaging
conditions as in (e) after moving the laser fiber-optic.
3. Point spread function (PSF) assessment
3.1 Rationale
A point spread function assessment of an objective lens will provide two pieces of information: 1) The quality of the
objective lens and 2) the resolution of the microscope optical system when imaging with that lens. All new objective
lenses should be tested before they are used routinely to ensure they have been well manufactured. Lenses should also
be tested on a regular basis or when image quality is compromised to assess if repairs are needed [1].
3.2 Methods
Briefly, a slide containing sub-resolution fluorescent microspheres (100 nm diameter for a 63x 1.4 NA oil immersion
lens) is imaged in 3D. Figure 2a and 2b show schematics of a point source laterally in the xy image plane and axially in
the yz image plane, respectively. Figure 2c and 2d show the resulting image of this point source laterally and axially.
The point source microspheres will appear much larger when imaged by the microscope due to distortions by the optics
owing mainly to the diffraction of light. The light is spread out in the resulting image so it is often termed the point
spread function (PSF). This phenomenon was first described by Sir Airy so the resulting image is also termed the Airy
pattern or the Airy disk. The PSF and Airy disk terms are often used interchangeably in microscopy. Laterally, in the xy
image plane the function appears as a spot with concentric diffraction rings around it, while axially the light is spread
out more extensively so the pattern is elongated and surrounded by a more complex pattern of diffraction. Inspection of
the PSF shape taken with a CLSM with the pinhole opened to ~5 Airy units can reveal a great deal about the objective
lens quality [1, 2]. A measurement of the full width half maximum (FWHM) of the PSF measured laterally and axially
with the pinhole set to 1 Airy unit can be compared to theoretical calculations. Doing so evaluates the resolution of the
microscope. The “Generate PSF report” function of the MetroloJ plug-in for FIJI (http://fiji.sc/wiki/index.php/Fiji ) can
be applied to 3D image stacks of single microspheres in order to measure the lateral and axial FWHM of the PSF. Note:
MeteroloJ overestimates the theoretical resolution so measured widths are typically larger than the reported theoretical
values[1].
3.3 Representative results
Figure 2e and 2f show high quality PSF images generated with a sample of 100 nm green microspheres using a 40x, 1.2
NA water immersion objective lens. MetroloJ analysis of this PSF data gives FWHM values of 0.28 μm (0.173 μm),
0.32 μm (0.173 μm) and 1.07 μm (0.506 μm) along the x, y and z axes, with the theoretical values from MetroloJ in
brackets.
3.4 Interpretation
A detailed interpretation of PSF quality can be found elsewhere [1-3]. Briefly, a good quality objective lens will have a
symmetric PSF whereas distortions in the PSF suggest problems with the optical light path, the objective lens itself or
other microscope components. See Table 1 in Cole et al. for high quality PSF images as well as images with artifacts
and their potential causes. Measurements from high quality, well-aligned microscope and objective lens are typically
within 40-60% of that predicted by theory. The theoretical values reported by MetroloJ can be multiplied by a factor of
1.25 to be more predictive of measured resolutions.
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3.5 Corrective measures
Details for corrective measures are given in Table 1 of Cole et al. [2]. These factors can be assessed and eliminated oneby-one followed by repeated measurements of the PSF. Some examples are to ensure DIC components are not in the
light path, that the objective lens and other optical elements are clean, that the CLSM pinhole is well-aligned and that
there is no thermal drift of the microscope during z-stack image acquisition. If these potential sources are rectified and
there are still issues with the PSF imaging and/or resolution then the objective lens should be inspected by the
microscope company to determine if it is in need of repair.
Fig. 2 Point source and theoretical lateral and axial PSFs
or Airy Disks. a) Schematic representation of a point source
in the lateral xy-axis. b) Schematic representation of a point
source in the axial z-axis. c) Theoretical lateral PSF
showing central Airy disk and diffraction rings. d)
Theoretical axial PSF showing central Airy disk and
diffraction rings. Reprinted with permission from Nature
Protocols [1].
4. Field Uniformity Assessment
4.1 Rationale
To obtain quantifiable intensity data, it is vital that the
illumination across the imaging field is uniform. This
test measures the homogeneity of light distribution
across the image field in three different directions:
horizontally across the center of the field of view and
along both diagonals from corner to corner. The
acceptable criteria are a maximum intensity variation
of 10% along the horizontal axis and 20% along the
diagonal axes [4].
4.2 Methods
The light source should be warmed up for at least one hour. Optical prisms and polarizers should be removed from the
optical path to avoid distortions in the resulting image. Figure 3a shows the placement of a fluorescent plastic slide on
the microscope stage. A 10x or 20x objective lens is put in place. Focus on the surface of the plastic slide. Figure 3b
shows an image of a scratch on the surface of the plastic slide that is used to aid in focusing. The microscope is then
focused ~20 µm into the slide. Figure 3c and 3d show images of the homogenous plastic slide in grey scale and with a
color look up table (LUT) for better visual contrast, respectively. For wide-field imaging, the camera exposure is set for
a mean gray level in the image of ~150 (8-bit image) or ~3500 (12-bit image). For a CLSM, the photomultiplier tube
(PMT) detector gain is set to achieve the same gray levels. 5.The PMT offset should also be set so all of the pixels
measure a positive intensity value. In either case, a high SNR is desirable, which may practically be obtained through
frame averaging of multiple images at low illumination intensity, to avoid photo bleaching of the plastic slide.
Draw a line across the horizontal or one of the diagonal (corner-to-corner) axes of the image. Measure the
fluorescence intensity along each line. Calculate the percent difference in intensity across each profile and compare the
values obtained to the accepted standards. Note that the overall image intensity is typically higher when the system is
well- aligned. If small-scale imperfections in the slide or high noise contributions affect the image uniformity, these
may be blurred out by applying a low-pass spatial filter or Gaussian kernel filter to the image before calculating the
percent variation for the purposes of the line-profile diagnostic.
4.3 Representative Results
A well-aligned microscope will exhibit a radially symmetric decrease in illumination about the center of the imaging
field. Figure 3c and 3d show data from a CLSM with a poorly aligned pinhole. From the data in Figure 3e, the system
does not pass the 10% variation criterion set for the horizontal intensity profile (solid red line), nor the diagonal
intensity profile criterion of less than 20% intensity variation (green dotted line and blue dashed line). Note that in this
figure, the normalization is set by the intensity profile maxima and not their means.
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4.4 Interpretation
If field uniformity is not within acceptable standards (10% variation along the horizontal, 20% variation along the
diagonal) but is symmetric then there may be an issue at the light source or with the coupling to the microscope. It can
also be due to sample not sitting flat in the slide holder on the microscope. If the illumination non-uniformity is
asymmetric then there is likely an issue with the alignment of the light source (lamp or laser) coupling to the
microscope or the detector alignment. On a wide-field microscope this could be due to a problem with the camera
alignment or camera-microscope coupling (wide-field, spinning disk confocal microscope, total internal reflection
fluorescence (TIRF) microscope). On a CLSM this could be the result of a problem with the laser coupling to the scan
head. Figure 3f shows a case where a transmitted light image is collected following Köhler alignment and reveals an
off-center illumination pattern. Figure 3g shows the same CLSM following re-alignment of the laser fiber coupling to
the confocal scan head. In other cases, the fluorescence confocal image such as those shown in Figure 3c and 3d reveal
that the pinhole is in need of re-alignment. In either case, the test should be repeated following re-alignment.
4.5 Corrective Measures
In certain cases, where intensity variations are symmetric, but exceed the above ranges, simply increasing the zoom
factor or moving to a higher resolution objective lens can result in intensity variations that are within range. For widefield microscopes, the most typical solution is to re-align the mercury bulb. The second thing to test is the lamp
coupling to the microscope. Camera alignment should be checked but is rarely the cause of non-uniform illumination.
For CLSMs, first verify that the pinhole is aligned. See the work of Cole et al. for details on aligning the pinhole [1].
Instrument manufactures can also perform system maintenance to check and re-align pinholes if necessary. If the laser
light source is not properly aligned, advanced users can use transmitted light imaging and align the laser fiber. Less
experienced users should contact their microscope service engineer.
It is always best to correct any microscope alignment issue at its source through hardware optimization. However, if
non-uniform field illumination still persists it can be corrected post image acquisition using standard images of uniform
samples such as fluorescent plastic slides. These corrections have been detailed elsewhere [5, 11]. Briefly, the specimen
is imaged and then an image of a uniform intensity sample (flatfield image) is collected with the same instrument
settings that were used to collect the raw image of the specimen. The raw specimen image (Imageraw) is multiplied by
the average intensity of the flatfield image (<IFF>). This value is then divided by the flatfield image (ImageFF). The
average background intensity in the corrected image is measured in an area where there is no fluorescence signal
(<IBG>) and this constant value is subtracted from each pixel in the image to give the fully corrected image (Imagecorr.).
=
[
∗
]
Equation 1
Fig. 3
Field Uniformity Tests. a) Orange
fluorescent test-slide on the microscope stage. b)
Image of the test-slide showing an in focus surface
scratch. c) Field uniformity image showing sample
line profiles for intensity profile assessments. d)
The same image as in 3c but enhancing the nonuniformity of illumination using a rainbow-colored
look-up table (LUT). e) Intensity profiles for lines
shown in 3c for a 440 nm laser. The green dotted
line and the blue broken line represent the diagonal
line profiles represented in 3c with the same colors.
The red line corresponds to the horizontal line
profile represented by a red line in 3c. The black
“x” line is the horizontal intensity profile in a wellaligned configuration f) Transmitted light image
with saturated pixels in red showing a misaligned
laser fiber to confocal head coupling. g)
Transmitted light image after fiber re-alignment.
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5. Light source stability
5.1 Rationale
For any quantitative light microscopy application it is critical that the excitation light source does not significantly vary
and introduce artifacts within the images. Tests should be performed on three different time scales. The short term
stability test monitors fluctuations in incident light intensity on the seconds to minutes time scale. This is relevant for
rapid time-lapse (e.g. calcium imaging) or 3D z-stack imaging where intensity variations would introduce artifacts in
the dataset. The moderate term stability test is on the three hour timescale which is relevant for a single user microscope
session where control and treated samples will be imaged and compared. Fluctuations in intensity or gradual increases
during light source warm-up could introduce artifacts when comparing sample images collected in a single imaging
session. Finally, the long term stability test is conducted over 300 hours and is relevant when comparing intensities of
samples between experimental imaging sessions. These tests will reveal any issues with hardware such as light source
stability, stability of laser powers (CLSM), issues with detectors or with confocal acousto-optic tunable filters (AOTF).
The acceptance criteria for these tests are an intensity deviation of 5% for the short term test and 10% for the moderate
and long term tests. These protocols are founded on those developed by Stack et al. [4] with some slight modifications
(e.g. power meter option).
5.2 Methods
The stability of the light source can be measured during warm-up to determine the optimal warm-up time for each light
source. Systems should be turned on and allowed to warm-up for the minimum required time before any performance
testing (e.g. 20-60 min). As with the field uniformity test, place a fluorescent plastic slide on the microscope stage
(Figure 3a). Use a 10x or 20x objective lens to focus on a scratch or a pen mark at the surface of the fluorescent slide
(Figure 3b). Then focus ~20 µm into the slide to achieve a homogenous area of the sample. For a wide-field microscope
set the camera exposure time for ~75% of the detector saturation (e.g. ~190 intensity units for an 8-bit image, ~3000
intensity units for a 12-bit image) and ensure that there are no saturated pixels. For a CLSM set the detector gain for
similar intensity values and ensure that the offset or black level is set so that no pixel measures zero intensity units.
Take an image of a small region of interest (ROI) of ~64x64 pixels in the center of the field of view. On the CLSM,
images can also be collected using the transmitted light photomultiplier tube detector to determine if alignment issues
are due to the pinhole or the laser fiber. Performing this type of measurement simultaneously on multiple lasers can rule
out issues with focus drift rather than laser power. For example, if the laser power is dropping for any of the lasers the
fluorescence images and transmitted light image stacks will follow exactly the same relative intensity patterns.
However, if it is a focus drift all of the fluorescence images will be affected in the same way but the transmitted light
measurements will not be drastically influenced. IMPORTANT: for the transmitted light measurements it is CRITICAL
that the microscope is in proper Köhler alignment.
•
•
•
Short Term Stability: Acquire an image every second for 5 minutes – total of 300 images.
Moderate Term Stability: Acquire an image every minute for 3 hours – total of 180 images.
Long Term Stability: Acquire an image every 10 hours for 300 hours – total of 30 images.
5.3 Representative Results
Images are analyzed by measuring the average intensity level of each frame and then calculating the % deviation, Dint,
over the entire series using Equation 2 where Imax is the frame with the highest average intensity reading, Imin the lowest
and Imean representing the average intensity over the entire time course. All of the images should be corrected for
background intensity prior to the calculation of the % deviation.
=
[
]
∗ 100
Equation 2
The acceptance criterion for this test is a maximum intensity deviation of 5% (short term) or 10% (moderate to long
term) [4]. Visual inspection of the images is also recommended in order to identify intensity fluctuations that may occur
within single images on a timescale of less than one second. This is important for CLSMs where it could reveal any
problems with rapid millisecond scale laser fluctuations. Figure 4a shows result for short term stability for a solid state,
mercury-free light source (blue curve), and two different mercury-based light sources (green and red curves). The
mercury-free light sources are an environmentally friendly alternative to mercury based sources and offer superior
stability (less than 1% deviation, the deviation is so small that it cannot be observed below the error bars for the
mercury light source #2). There are several publications outlining microscope light sources and the benefits of mercuryfree alternatives [12-15]. Figure 3b shows moderate term wide-field light source stability and Figure 3c long term
stability. In all cases, the mercury-free, solid state sources show superior performance.
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5.4 Interpretation
On the time-scale of the short term stability test, factors that typically contribute to sub-standard performance include
instability of lamps or lasers, intensity modulating filters and/or detectors [4]. Figure 4d shows that if instabilities are a
result of focal drift during testing of lasers on the CLSM then all laser power curves will drift with a similar % intensity
drop over time (blue and red curves). Transmitted light curves may show some drop in intensity as well but this
measurement is much less sensitive to focal drift than the fluorescence intensity measurement (green and purple
curves). Figure 4e shows an example of a laser that needs to be replaced as the power is oscillating with 10% intensity
changes over time (blue curve). This kind of serious artifact is exactly what this test is designed to find and would be
very difficult for an end-user to realize during normal operation of the microscope. This laser was replaced and the
power was very stable following the repair (green curve). The third curve in Figure 4e demonstrates the need to warmup lasers as they can take up to an hour to stabilize (red curve). To trouble shoot possible AOTF issues on a CLSM,
tests should be repeated with the laser light at 100%, essentially bypassing the AOTF function. If problems persist it is
unlikely that the AOTF is the issue.
5.5 Corrective Measures
Typically problems with intensity stability suggest lamp or laser stability issues. Firstly, potential sources such as stage
drift or focal drift should be ruled out. For mercury lamp based systems, bulb replacement and/or alignment is/are likely
required. For laser-based systems, the pinhole alignment should be verified and then the laser power supply and/or the
laser itself. It is also possible that the AOTF may need to be replaced. The assistance of a field service specialist is
likely required in order to repair or replace these components. Focal drift can be corrected by reducing air currents
around the microscope and leaving sufficient time for the system to warm-up. Proper warm-up times for lamps and
lasers should abate fluctuations in the power of a light source that is performing well.
6. CLSM laser power linearity and power meter measurements.
6.1 Rational
Using the tests in Section 5 it is often difficult to determine if CLSM laser power issues are due to the functioning of the
AOTF. Measurements of laser power at different AOTF settings is a way of testing if the filter is responding and
calibrated properly. Monthly absolute power measurements are a proactive way of identifying when any light source
needs to be replaced or repaired. For single time-point measurements it is often much more straightforward to make the
measurements using a power meter rather than a fluorescent plastic slide.
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Fig. 4 Long term laser stability results. a) Short-term stability tests of mercury free solid state light source and two mercury based
wide-field microscope sources. b) Moderate term stability tests of the same light sources as in a. c) Long term stability test of the
same light sources as in a. d) Moderate term stability of a 473 nm, and a 594 nm laser on a CLSM measured during the same time
course. Intensities were measure from fluorescence images as well as transmitted light images. The drift of both intensity levels
simultaneously points towards a likely focal drift during the measurement. e) Moderate term stability of a 561 nm laser before (blue
line) and after (green line) being replaced. Laser warm-up of a 488 nm laser line (red line). Error bars are standard deviation.
6.2 Methods
Warm-up the lasers for at least 60 minutes and place a low magnification (e.g. 10x/0.3 NA) objective lens in place.
Figure 5a shows a suitable power meter and Figure 5b shows the power meter sensor directly on top of the objective
lens. It can be secured in place with the stage sample holder or sticky-tack. Set the power meter to the selected
wavelength to be measured (e.g. 488 nm) and zero the instrument with low light levels in the room. Record laser powers
with the AOTF to 20%, 40%, 60%, 80% and 100% output. Repeat the process for all dichroic mirror/laser combinations
to be tested. To uncouple AOTF and laser difficulties it is best to measure the actual power at the output from the laser
with no optical components in the light path. Of course safety goggles and proper laser safety procedures should be
followed when working with the laser output from the source. In addition, by plotting the measured laser power
readings against the corresponding estimated values, one can obtain a rough approximation of the transmission
efficiency (TE) of the beam-path from the slope of the curve.
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6.3 Representative Results
Figure 5c shows laser power measurements of a 633 nm laser line with a poorly calibrated AOTF (dots and solid line).
Following recalibration by a field service specialist a proper linear calibration was achieved (triangles and dotted line).
Figure 5d shows single laser power measurements at 100% power at the 10x objective lens measured several months
apart. Two of the lasers have relatively stable power measurements, while one (dotted line) shows a marked power
reduction at time-point 3. This kind of power drop can be predictive of imminent laser failure. However, these types of
measurements should be repeated to rule out user errors, such as poor zeroing of the sensor or poor positioning of the
sensor on the objective before the laser is repaired. For instance, if the laser is not hitting the sensor exactly normal there
can be intensity errors on the order of 20%.
6.4 Interpretation
Non-linearity of the AOTF is almost always due to improper calibrations. Dips in laser power are often due to poor
measurements and should be repeated several times before seeking laser repair. Consistent decreases in laser power
over time suggest imminent laser failure. The laser should be replaced before a complete failure occurs.
Fig. 5 a) A Coherent FieldMaxII-TO laser power meter. b) Visible light sensor mounted onto the microscope stage with a 10x
objective lens. It is held in place by the stage insert. c) Following the installation of a new AOTF controller, the transmission profile
of the 633 nm laser line of an LSM510 confocal microscope displayed a marked degree of non-linearity (solid line). After the field
service specialist recalibrated the AOTF, a linear efficiency function was restored (dashed line). The maximum laser power at the
10x objective lens of the 488 nm laser line on several different CLSMs plotted over successive time points.
6.5 Corrective Measures
AOTF calibrations and power supply or laser repairs or replacements are typically conducted by a service engineer.
Have your data ready to show to service or sales representatives in order to demonstrate that the instrument
performance is sub-optimal. While the methods and examples provided relate to the CLSM these tests can easiliy be
adapted to test any fluorescence microscope light source.
7. Co-registration Test
7.1 Rationale
When imaging multiple fluorescent probes, a high fidelity in the registration between the various detection channels is
essential to produce multicolor images that are accurately superimposed. This is even more crucial when the intention is
to obtain meaningful co-localization information. This test measures the co-registration of the center-of-mass along the
x, y and z-axis of a multi-color microsphere image acquired in multiple detection channels. The co-registration should
be within 1 pixel in the lateral and axial directions to be deemed “acceptable” [4].
7.2 Methods
Ensure the pinholes on a CLSM are well-aligned before conducting this test. Image 4 µm multi-colored microspheres
(Tetraspeck™ by Molecular Probes, Cat. #T7283) using a high resolution (e.g. 63x/1.4 NA) immersion objective lens.
Imaging parameters should be set up as described in Section 3. Ensure to sample with Nyquist criteria or higher [1] with
settings such as a digital zoom of 5-10x and a z-stack with an image step-size of 0.1 µm. Centre a microsphere in the
field of view using the stage movement to ensure the microsphere is at the center of the objective lens for minimal
aberrations. Collect z-stacks of images for 2-4 of the relevant colors of the multi-colored microspheres simultaneously.
Set up the filters and detectors in each channel in the same way they will be set during future experimentation.
Microsphere image stacks are then analyzed using the MetroloJ plugin “Co-alignment Report” function in FIJI [2, 4].
Up to three channels can be simultaneously assessed in the report. To obtain accurate dimensional calculations, the
operator must input the magnification and NA of the objective lens as well as the peak emission wavelengths being
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measured in each channel. In addition to generating xy, xz and yz profiles of the image stack for visual assessment, the
report generates table matrices describing the relative center-of- mass positions of each channel in pixels and/or µm.
7.3 Representative results
Figures 6a and 6b show results from two poorly-aligned imaging channels with a large pixel shift between the locations
of the microsphere imaged in the two channels. Figures 6c and 6d show results of two aligned image channels. Similar
measurements need to be done along the z-axis of the image stack.
7.4 Interpretation of the data
While lateral (xy) alignment shifts can often be attributed to laser or confocal pinhole misalignment, the relative image
registration of the optical components between alternative beam path configurations including different dichroic mirrors
also plays a critical role. Additional sources of poor co-registration include poor quality lenses, the presence of
chromatic aberrations and incorrect laser collimator positioning. The further apart the wavelengths of the two colors are
more difficult to achieve high quality co-registration. Z-axis alignment is not shown here but can be particularly
problematic with large wavelengths and/or poorly corrected optics [16].
Fig. 6 a) Image of a Tetraspeck™ microsphere in two CLSM channels with poor alignment of the blue fluorescence (cyan, 405 nm
excitation, 420 nm longpass emission) and green fluorescence (red, 488 nm excitation, 505-530 nm emission). b) Intensity profile
across the microsphere images shown in a. c) Image of a Tetraspeck™ microsphere in the same channels as a, but after alignment of
the laser fiber coupling to the scan head. d) Intensity profile across the microsphere images shown in c.
7.5 Corrective measures
Ensure good alignment of all pinholes in the system with the exact optics in place as will be used for imaging. Ensure
DIC optics are not present in the light path (they can result in double images). Although sensitivity will be reduced it is
best to use multi-dichroic for multiple colors imaging when possible to avoid slight errors in co-registration between
different dichroic mirrors. If high sensitivity is important (e.g. live cell or organism imaging) carefully measure any
shifts between channels in x, y, and z and correct image stacks post acquisition. When possible, use dyes that are not too
spectrally separated. If using blue and far red dyes together be sure to measure and correct for shifts in co-registration,
which will be particularly apparent along the z-axis [16]. For wide-field microscopes, use high quality fluorescent cubes
that are factory-calibrated to have negligible registration shifts relative to each other, often called “shift-free”. Test these
fluorescence cubes when the microscope is purchased and periodically to ensure optimal performance.
8. Conclusion
The main goal of this chapter is to present a practical guide to ensure the proper care, effective maintenance and quality
control of light microscopy equipment. The need for proper maintenance and operation of the light microscope to
ensure the highest quality artifact-free data for scientific studies cannot be understated. It is our hope that this review
will assist microscopists at all levels in maintaining their microscopes. We continue to refine and develop our protocols
and collaborate with others to develop new methods to benchmark fluorescence microscopy (www.abrf.org/lmrg;
www.nist.gov/mml/bbd/cell_systems/fluorescence-microscope-benchmarking.cfm).
Acknowledgements We acknowledge the work of Richard Cole and the Association of Biomolecular Resource Facilities (ABRF)
Light Microscopy Research Group for their work on light microscopy quality control which is the bases for a lot of this work and has
inspired us to ensure our instrumentation is performing at its peak. Thank you to David Verbich and Judith Lacoste who provided
images for Figure 1. Thank you to Richard Cole, Judith Lacoste, Thoma Kareco and Denis Filipo for critical discussions of
microscope maintenance and for sharing their wealth of technical expertise. Funding for this project and all quality control data was
collected at the McGill University Life Sciences Complex Advanced BioImaging Facility (ABIF).
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