microscope maintenance and quality control: a practical · pdf filemicroscope maintenance and...

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), 2 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

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 co-registration 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; co-localization; 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. Ammonia-free 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



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 one-by-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.



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 wide-field 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 non-uniformity 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 well-aligned 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.


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 mercury-free 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.



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 warm-up 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.


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.




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.


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).



References

[1] Cole RW, Jinadasa T, Brown CM. Measuring and interpreting point spread functions to determine confocal microscope resolution and ensure quality control. Nature protocols. 2011; 6:1929-41.

[2] Cole RW, Thibault M, Bayles CJ, Eason B, Girard A-M, Jinadasa T, Opansky C, Schulz K, Brown CM. International Test Results for Objective Lens Quality, Resolution, Spectral Accuracy and Spectral Separation for Confocal Laser Scanning Microscopes. Microscopy and Microanalysis. 2013; FirstView:1-16.

[3] Goodwin PC. Evaluating Optical Aberration Using Fluorescent Microspheres: Methods, Analysis, and Corrective Actions. Digital Microscopy, 3rd Edition. Sluder GWolf DE. 2007; 81:397-413.

[4] Stack RF, Bayles CJ, Girard AM, Martin K, Opansky C, Schulz K, Cole RW. Quality assurance testing for modern optical imaging systems. Microscopy and microanalysis : the official journal of Microscopy Society of America, Microbeam Analysis Society, Microscopical Society of Canada. 2011; 17:598-606.

[5] Lacoste J, Young K, Brown CM. Live-cell migration and adhesion turnover assays. Methods in molecular biology. 2013; 931:61-84.

[6] Zucker RM, Price O. Evaluation of confocal microscopy system performance. Cytometry. 2001; 44:273-94. [7] Webb RH. Confocal Optical Microscopy. Rep. Prog. Phys. 1996; 59:427-471. [8] Zucker RM, Price OT. Practical confocal microscopy and the evaluation of system performance. Methods. 1999; 18:447-58. [9] Lacoste J, Vining C, Zuo D, Spurmanis A, Brown CM. Optimal Conditions for Live Cell Microscopy and Raster Image

Correlation Spectroscopy (RICS). Annual Reviews in Fluorescence 2010. Geddes CD. 2011; [10] Frigault MM, Lacoste J, Swift JL, Brown CM. Live-cell microscopy - tips and tools. Journal of cell science. 2009; 122:753-67. [11] Webb DJ, Brown CM. Epi-fluorescence microscopy. Methods in molecular biology. 2013; 931:29-59. [12] Wessels JT, Pliquett U, Wouters FS. Light-emitting diodes in modern microscopy--from David to Goliath? Cytometry A. 2012;

81:188-97. [13] Cole RW, Turner JN. Light-emitting diodes are better illumination sources for biological microscopy than conventional

sources. Microscopy and microanalysis : the official journal of Microscopy Society of America, Microbeam Analysis Society, Microscopical Society of Canada. 2008; 14:243-50.

[14] Biard R, Kaufman D, Brown CM. Mercury Free Microscpy: An Opportunity for Core Facility Directors. J. Biomol Tech. 2014; July:In Press.

[15] Aswani K, Jinadasa T, Brown CM. Fluorescence Microscopy Light Sources. Microscopy Today. 2012; 20:22-28. [16] North AJ. Seeing is believing? A beginners' guide to practical pitfalls in image acquisition. J Cell Biol. 2006; 172:9-18.


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