Interview Questions for Microscopy Instrumentation

Light microscopy relies on the interaction of visible light with the specimen. A light source illuminates the sample, and the light that passes through or is reflected from the sample is then magnified by a series of lenses. The resulting image is projected onto the viewer’s eye or a detector. Think of it like using a magnifying glass, but with much higher magnification and resolution achieved through multiple lenses and sophisticated illumination techniques.

The key principle is the bending (refraction) of light as it passes through different mediums. Lenses are precisely shaped to control this refraction, focusing the light to create a magnified image. The resolution – the ability to distinguish between two closely spaced points – is limited by the wavelength of light. This means there’s a fundamental limit to how small an object we can see clearly with light microscopy.

These microscopy techniques differ primarily in how they manipulate the light to enhance contrast and visibility of the specimen:

• Brightfield Microscopy: This is the most common type. The sample is illuminated directly from below, and the image is formed by the light that passes *through* the sample. Stained samples are usually required to enhance contrast, as unstained samples often appear transparent. Imagine shining a flashlight through a translucent object – you’ll only see details if there’s sufficient contrast.
• Darkfield Microscopy: In this method, only the light scattered by the sample reaches the objective lens. This results in a bright specimen against a dark background. This is particularly useful for observing unstained, transparent specimens, as it highlights the edges and boundaries of the sample. Think of it like seeing dust motes illuminated in a darkened room; you don’t see the air itself, only the scattered light from the dust.
• Phase-Contrast Microscopy: This technique converts differences in refractive index within the sample into visible contrast. It’s particularly useful for examining living cells and other unstained transparent specimens, allowing visualization of internal structures without the need for staining. This method essentially takes advantage of the subtle variations in how light passes through different parts of the cell, transforming those variations into visible changes in brightness.

Fluorescence microscopy uses fluorescent molecules (fluorophores) to label specific structures within a sample. These fluorophores absorb light at a specific wavelength (excitation wavelength) and then emit light at a longer wavelength (emission wavelength). This emitted light is then detected to create an image.

• Advantages: High specificity (labeling specific structures), high sensitivity (detecting small amounts of target molecules), multicolor imaging (labeling multiple structures simultaneously), and allows for studying dynamic processes in living cells.
• Disadvantages: Photobleaching (fluorophores lose their fluorescence over time), phototoxicity (fluorophores can damage cells), cost of equipment and fluorophores, and requires specialized sample preparation.

For example, in biomedical research, fluorescence microscopy is crucial for visualizing the location and distribution of proteins within cells, tracking the movement of molecules, and studying cellular interactions.

Confocal microscopy eliminates out-of-focus light, resulting in sharper, higher-resolution images. It achieves this by using a pinhole aperture in front of the detector. A laser scans the sample point by point, and only the light emitted from the focal plane passes through the pinhole. This creates optical sections, which can be stacked to construct a 3D image of the sample.

Imagine taking a stack of very thin slices of bread. Each slice represents an optical section. By carefully arranging these slices, you reconstruct the entire loaf of bread, analogous to creating a 3D image from the optical sections in confocal microscopy. This technique is particularly important in visualizing thick samples or samples with complex three-dimensional structures.

The fundamental difference lies in the type of radiation used to image the sample. Light microscopy uses visible light, while electron microscopy uses a beam of electrons. This difference leads to significant variations in resolution and magnification. Electron microscopy provides significantly higher resolution (down to the nanometer scale) compared to light microscopy (limited by the wavelength of light), allowing for visualization of much smaller structures. This is because electrons have a much shorter wavelength than visible light.

Another key difference is sample preparation. Electron microscopy requires complex sample preparation techniques, often involving fixation, dehydration, and embedding in resin to create ultrathin sections for transmission electron microscopy (TEM).

There are several types of electron microscopy, each with its own applications:

• Transmission Electron Microscopy (TEM): Electrons pass *through* a very thin sample, creating an image based on the electron density of different parts of the sample. TEM provides high resolution and is excellent for visualizing internal structures of cells and materials.
• Scanning Electron Microscopy (SEM): A focused beam of electrons scans the surface of the sample, and the emitted electrons or X-rays are detected to create a three-dimensional image of the surface topography. SEM provides detailed information about surface morphology and is widely used in materials science and biology.
• Scanning Transmission Electron Microscopy (STEM): This combines aspects of TEM and SEM, offering both high-resolution imaging of internal structures and surface information. STEM often uses specialized detectors for advanced analytical capabilities.

Sample preparation for electron microscopy is crucial for obtaining high-quality images. It is a multi-step process and depends on the type of microscopy (TEM or SEM) and the nature of the sample:

• Fixation: Preserves the sample’s structure by cross-linking proteins. This prevents degradation and distortion.
• Dehydration: Removes water from the sample, preventing damage during embedding.
• Embedding: The sample is embedded in a resin to provide support during sectioning.
• Sectioning (for TEM): The embedded sample is sliced into ultrathin sections (typically 50-100 nm thick) using an ultramicrotome.
• Staining (often for TEM): Heavy metal stains (e.g., uranyl acetate, lead citrate) are used to increase contrast and reveal fine details.
• Coating (often for SEM): A conductive coating (e.g., gold, platinum) is applied to the sample surface to prevent charging during SEM imaging.

The specific procedures vary greatly depending on the sample type, but the goal is always to maintain the sample’s structural integrity and enhance visibility for imaging.

Resolution in microscopy refers to the ability of a microscope to distinguish between two closely spaced objects as separate entities. Think of it like the sharpness of an image – a higher resolution means you can see finer details. It’s determined by the smallest distance between two points that can still be perceived as distinct. Poor resolution results in blurry, indistinct images, while high resolution provides crisp, clear images revealing intricate structures.

The limit of resolution is fundamentally constrained by the diffraction of light. This means light waves bend around objects, blurring the image. The Rayleigh criterion is a commonly used formula to define this limit: d = 0.61λ / NA, where ‘d’ is the minimum resolvable distance, ‘λ’ is the wavelength of light, and ‘NA’ is the numerical aperture (explained below). This shows that shorter wavelengths and higher numerical apertures lead to better resolution.

For example, a high-resolution electron microscope can resolve features much smaller than an optical microscope because it uses electrons with much shorter wavelengths than visible light.

Numerical aperture (NA) is a crucial measure of a microscope objective’s ability to gather light and resolve fine details. It’s a dimensionless number that represents the light-gathering power of the objective lens. A higher NA indicates a greater ability to gather light and, consequently, better resolution and image brightness. It’s calculated using the formula: NA = n sinθ, where ‘n’ is the refractive index of the medium between the objective lens and the specimen (usually air or immersion oil), and ‘θ’ is half the angle of the cone of light entering the objective lens.

The significance of NA lies in its direct impact on resolution, as seen in the Rayleigh criterion. Using immersion oil with a high refractive index (n) increases the NA, allowing for a much smaller resolvable distance ‘d’ and thus better resolution. Imagine trying to see small details underwater; the water acts like immersion oil, helping to bring those details into focus.

Magnification and resolution are distinct but related concepts in microscopy. Magnification refers to the increase in the apparent size of an object, while resolution refers to the ability to distinguish between two closely spaced objects. You can magnify an image many times, but if the resolution is poor, the magnified image will still be blurry and lack detail. High magnification without sufficient resolution is essentially useless, producing a large, blurry image.

Think of it like zooming in on a photograph. You can zoom in (magnify) to make the picture larger, but if the original photo was taken with a low-resolution camera, zooming will only enlarge the pixels, making the image pixelated and blurry. A high-resolution photograph will allow you to zoom in and still maintain sharp detail.

Therefore, high resolution is essential for meaningful magnification; it’s pointless to magnify something you can’t clearly resolve.

Depth of field (DOF) in microscopy refers to the range of distances along the optical axis of the microscope where the specimen appears to be in sharp focus. A large depth of field means a greater range of distances is in focus, while a small depth of field means only a very thin slice of the specimen will be sharply focused. This is directly related to the aperture of the objective lens; smaller apertures result in a larger depth of field and vice versa.

Imagine taking a photograph of a flower. A large DOF would keep both the petals in the foreground and the background leaves in sharp focus, while a small DOF would only keep one sharply in focus, blurring the rest. In microscopy, this can be critical when imaging thick samples. A smaller depth of field, for instance, is crucial in confocal microscopy to ensure sharp images of specific depths in a thick sample.

The DOF is affected by several factors, including the magnification of the objective lens, the numerical aperture, and the wavelength of light. Generally, higher magnification and higher NA mean smaller DOF.

Microscope objectives are the most critical components, determining the image quality. Several types exist, each with specific characteristics:

• Achromatic objectives: Correct for chromatic aberration (color fringing) for two wavelengths (usually red and blue) and spherical aberration (blurring due to lens shape) for one wavelength.
• Apochromatic objectives: Correct for chromatic aberration for three or more wavelengths and spherical aberration for two or more wavelengths. They provide superior image quality, but are more expensive.
• Plan objectives: Correct for field curvature, ensuring a flat field of view, meaning the entire field is in focus, not just the center. This is crucial for imaging large samples.
• Plan-apochromatic objectives: Combine the benefits of plan and apochromatic objectives, providing the highest level of correction for aberrations and a flat field of view.
• Oil immersion objectives: Use immersion oil between the objective and the specimen to increase the numerical aperture and resolution. The oil has a refractive index similar to glass, reducing light refraction and improving the light-gathering ability of the objective.
• Water immersion objectives: Similar to oil immersion, but use water as the immersion medium. They are often used for live cell imaging as water is less damaging to cells.

The choice of objective depends on the specific application and desired image quality. Higher-quality objectives (apochromatic, plan-apochromatic) offer superior performance but are significantly more expensive.

Microscopy images can be affected by various artifacts that can obscure or misrepresent the true structure of the specimen. Common artifacts include:

• Spherical aberration: Blurring due to imperfect lens curvature.
• Chromatic aberration: Color fringing due to different wavelengths of light being refracted differently.
• Diffraction: Bending of light waves around objects, resulting in blurred edges.
• Halos: Bright rings around objects.
• Streaks: Linear artifacts.
• Noise: Random variations in pixel intensity.

Minimizing these artifacts requires careful attention to technique:

• Using high-quality objectives: Apochromatic and plan-apochromatic objectives minimize aberrations.
• Proper sample preparation: Avoiding bubbles, debris, and uneven staining.
• Careful focusing: Achieving the correct focal plane minimizes diffraction and other artifacts.
• Appropriate illumination: Using Köhler illumination to achieve uniform illumination.
• Image processing: Removing noise and other artifacts using image analysis software.

Understanding the causes of artifacts is essential for interpreting microscopy images accurately. A poorly prepared sample or incorrectly used microscope can easily introduce artifacts leading to misinterpretations.

Microscope calibration ensures accurate measurements and reliable data. The process involves verifying the accuracy of magnification and stage movement. Calibration typically uses a stage micrometer, a slide with a precisely etched scale (e.g., 1 mm divided into 100 or 1000 μm). The procedure differs slightly depending on the microscope type, but the general steps are:

1. Prepare the stage micrometer: Place the stage micrometer on the microscope stage.
2. Select a high-power objective: Use an objective lens with a known magnification.
3. Focus on the micrometer scale: Adjust the focus to clearly visualize the scale.
4. Measure a known distance: Use the eyepiece reticle (or an ocular micrometer, a scale within the eyepiece) to measure a known distance on the stage micrometer (e.g., 1 mm). Record the number of divisions on the eyepiece reticle that correspond to this known distance.
5. Calculate the calibration factor: This factor converts eyepiece reticle divisions to real-world units. For example, if 10 eyepiece divisions correspond to 1 mm (1000 μm), the calibration factor is 100 μm/division.
6. Repeat with other objectives: Repeat steps 3-5 for each objective lens to determine the calibration factor for each magnification.

Regular calibration is crucial for accurate measurements. It is usually done before and after each experiment or at regular intervals (depending on use) to ensure that measurements obtained are dependable and comparable across different experiments or sessions.

Image acquisition in microscopy involves capturing the light or electrons interacting with your sample to form an image. This process starts with illuminating the sample using a light source (for light microscopy) or an electron beam (for electron microscopy). The light or electrons that pass through or are reflected from the sample are then collected and focused by a series of lenses to create a magnified image. This image is then captured by a detector, often a camera, converting the light or electron signal into a digital image file.

Image processing refines this raw image to enhance its quality and extract meaningful information. This can involve various techniques such as adjusting brightness and contrast, removing noise, sharpening the image, and applying filters to highlight specific features. For example, deconvolution algorithms can computationally remove blurring caused by the microscope’s optics, resulting in a sharper, higher-resolution image. Software packages like ImageJ or Fiji are commonly used for image processing, offering a wide array of tools and plugins for advanced image analysis.

Imagine taking a photo of a flower. Acquisition is like taking the picture; processing is like editing it in Photoshop to improve brightness, contrast, and remove blemishes, revealing more detail.

Microscopy image analysis employs a variety of techniques to quantify and interpret the visual information. These techniques can be broadly categorized as:

• Quantitative analysis: This involves measuring features within the image, such as the size, shape, intensity, and number of objects. For instance, we might measure the area of cells in a tissue sample or count the number of bacteria in a culture. Software like ImageJ provides tools for automated measurements.
• Qualitative analysis: This involves visual inspection of the image to identify and characterize features. For example, examining the morphology of cells to identify different cell types or observing the distribution of specific proteins within a cell.
• Morphometric analysis: This deals with the measurement of shape and size of structures within the image. This might involve measuring the length and width of filaments or the perimeter and area of cells, to assess their size and shape changes.
• 3D reconstruction: When using techniques like confocal microscopy or electron tomography, multiple images are acquired at different focal planes to build a three-dimensional representation of the sample. This allows for a much deeper understanding of the sample’s structure.

The choice of technique depends heavily on the research question and the type of microscopy used. For example, analyzing the fluorescence intensity of specific proteins requires different tools than measuring the size distribution of particles in a material sample.

Troubleshooting microscopy problems requires a systematic approach. It’s like detective work! Here’s a general framework:

1. Check the obvious: Is the microscope turned on? Are the light source and appropriate filters in place? Is the sample correctly mounted and in focus?
2. Assess the image quality: Is the image blurry, too dark, too bright, or noisy? This helps narrow down the problem area.
3. Examine the optics: Clean the lenses with appropriate lens paper and cleaning solution. Check for any misalignments or damage to the lenses. Dirt on the lenses is a very common cause of poor image quality.
4. Verify the illumination: Is the light source properly adjusted? Are the condenser and aperture correctly aligned for optimal illumination? A properly adjusted Köhler illumination is crucial for optimal image quality in light microscopy.
5. Evaluate the sample: Is the sample properly prepared and stained? Is the sample thick or too dense, preventing light from passing through (in transmission microscopy)?
6. Check the camera settings: If using a digital camera, ensure proper exposure, gain, and white balance settings.
7. Consult the microscope manual: This invaluable resource contains troubleshooting guides specific to your microscope model.

If the problem persists, seek assistance from experienced microscopists or the manufacturer.

Proper sample mounting is crucial for obtaining high-quality microscopy images. It ensures the sample is stable, correctly positioned, and doesn’t move during image acquisition. Inappropriate mounting can lead to blurry images, artifacts, and inaccurate measurements. The method depends on the type of sample and microscopy technique used.

• Stability: The mount must securely hold the sample, preventing drift or movement. This is especially important for high-magnification imaging or time-lapse experiments.
• Correct Orientation: The sample should be oriented appropriately for optimal visualization of the features of interest.
• Minimizing Artifacts: The mounting medium should be compatible with the sample and the imaging technique to avoid introducing artifacts or distortions.
• Accessibility: The sample should be easily accessible for observation and manipulation under the microscope.

For example, a thin tissue section needs to be mounted on a glass slide with a mounting medium that preserves its structure, whereas a live cell culture might require a special environmental chamber to maintain its physiological conditions.

Staining techniques are crucial for visualizing specific structures or components within a sample. They enhance contrast and reveal details that might be otherwise invisible. Many staining techniques exist, categorized by their application:

• Hematoxylin and Eosin (H&E): A standard histological stain used in pathology to visualize cell nuclei (hematoxylin, purple) and cytoplasm (eosin, pink). This provides a general overview of tissue morphology.
• Gram staining: Used in microbiology to differentiate bacteria into Gram-positive (purple) and Gram-negative (pink) based on their cell wall composition.
• Immunofluorescence: Uses fluorescently labeled antibodies to target specific proteins or antigens. Different fluorophores can be used to label multiple targets simultaneously.
• DAPI staining: A fluorescent dye that binds to DNA, enabling visualization of cell nuclei.
• Periodic acid-Schiff (PAS) staining: Detects polysaccharides and glycogen in tissues, staining them magenta or purple.

The choice of stain depends on the target and the desired information. For instance, if one wishes to study the distribution of a specific protein within a cell, immunofluorescence would be appropriate, whereas if one wants a general overview of tissue architecture, H&E staining is a common choice.

Microscope maintenance is essential for optimal performance and longevity. Regular cleaning and careful handling are crucial.

• Daily cleaning: Gently wipe down the microscope body and stage with a soft, lint-free cloth. Remove any dust or debris using compressed air.
• Lens cleaning: Clean the objective lenses regularly with specialized lens paper and lens cleaning solution. Use a circular motion to avoid scratching the lens surface. Always start with the lowest magnification objective.
• Periodic maintenance: Schedule regular professional servicing by a qualified technician. This involves more thorough cleaning, alignment checks, and potential repairs.
• Proper storage: Store the microscope in a clean, dry environment, covered with a dust cover to protect it from dust and environmental contaminants.

Neglecting maintenance can lead to poor image quality, damage to the instrument, and ultimately, costly repairs. Regularly cleaning the lenses and ensuring a dust-free environment is crucial. Think of it as regular car maintenance; it keeps the machine running smoothly.

Safety precautions are paramount when using a microscope. Here are some key considerations:

• Eye safety: Never look directly into the light source. Use appropriate filters to protect your eyes from harmful radiation.
• Handling caution: Handle the microscope carefully to avoid dropping or damaging it. Use both hands when moving the microscope to ensure stability.
• Electrical safety: Ensure the microscope is properly grounded to prevent electric shock. Never use the microscope near water or other liquids.
• Sample handling: Use appropriate personal protective equipment (PPE) when handling samples, especially if they contain hazardous materials.
• Ergonomics: Maintain a proper posture while using the microscope to prevent fatigue and strain.
• Chemical safety: When using staining solutions or mounting media, always wear appropriate PPE, work in a well-ventilated area, and dispose of waste according to safety regulations.

Prioritizing safety not only protects your health but also ensures the longevity and proper functioning of your microscope. Following these guidelines is crucial for responsible microscope usage.

My experience with microscopy software spans a wide range of platforms, from basic image viewers to advanced image processing and analysis suites. I’m proficient in using software such as ImageJ/Fiji (including plugins like 3D Viewer, CellProfiler, and TrakEM2), NIS-Elements (Nikon), ZEN (Zeiss), and Huygens Professional (Scientific Volume Imaging). Each software package offers unique functionalities tailored to specific microscopy techniques and experimental designs. For instance, ImageJ/Fiji’s open-source nature and extensive plugin library make it incredibly versatile for image manipulation, measurement, and analysis across diverse applications. NIS-Elements is particularly powerful for advanced microscopy techniques like confocal microscopy, offering robust features for image acquisition and deconvolution. I’ve used these programs extensively for tasks like image stitching, noise reduction, 3D reconstruction, colocalization analysis, and quantitative measurements of fluorescence intensity.

For example, in a recent project involving live-cell imaging, I utilized NIS-Elements to acquire time-lapse images of cell migration. Subsequently, I employed ImageJ/Fiji to track individual cell movements and analyze their speed and trajectory, using the Manual Tracking plugin. This involved carefully adjusting parameters such as thresholding to accurately identify cells across multiple frames.

I have extensive experience with various advanced microscopy techniques, including immunofluorescence (IF), in situ hybridization (ISH), and confocal microscopy. Immunofluorescence involves labeling specific cellular components or proteins using fluorescently tagged antibodies. I routinely perform IF staining, optimizing protocols for various antibody combinations and tissue types to minimize background noise and achieve high signal-to-noise ratios. This often involves troubleshooting issues like non-specific binding and antibody cross-reactivity.

In situ hybridization (ISH) allows visualization of specific nucleic acid sequences within cells or tissues. My expertise includes both fluorescent ISH and chromogenic ISH, utilizing techniques such as RNA-ISH and FISH (fluorescence in situ hybridization). I have experience optimizing ISH protocols for different target sequences and tissue types, ensuring specificity and sensitivity of the assay. For example, I’ve worked on projects using FISH to identify and quantify specific chromosomal abnormalities in cancer cells.

Confocal microscopy allows for 3D imaging with high resolution and minimal background noise. I’m proficient in operating confocal microscopes and analyzing the acquired images, including using deconvolution algorithms to improve image quality and 3D rendering for visualization of complex structures.

Quality control of microscopy images is paramount to ensure the reliability and validity of experimental findings. My approach involves a multi-step process starting even before image acquisition. This begins with meticulous sample preparation, including optimizing fixation and staining protocols. During image acquisition, parameters such as laser power, gain, and exposure time are carefully controlled and documented to avoid artifacts or photobleaching. Post-acquisition, a rigorous quality assessment is performed.

This assessment involves checking for artifacts like uneven illumination, noise, and out-of-focus blur. For example, I routinely examine images for the presence of autofluorescence, a common issue which can be mitigated by appropriate sample preparation techniques and image processing. I also assess the image quality using metrics such as signal-to-noise ratio and resolution. Any potential issues are flagged and addressed through image processing techniques like background subtraction, noise reduction, and deconvolution. Finally, appropriate scaling, labeling, and metadata are incorporated for clear presentation and reproducibility.

Data analysis and interpretation from microscopy experiments are crucial for drawing meaningful conclusions. My experience encompasses a range of quantitative image analysis techniques. I’m proficient in using image processing software to quantify fluorescence intensity, measure cell sizes and shapes, and analyze colocalization patterns. For instance, I often use ImageJ/Fiji’s measurement tools to quantify the expression levels of different proteins in individual cells, using the ‘Analyze Particles’ function to characterize features of interest within an image.

Beyond basic measurements, I utilize more advanced analysis methods, including statistical analysis, to test hypotheses and draw inferences from the experimental data. I’m familiar with statistical software such as R and Python, using packages like ‘ggplot2’ and ‘SciPy’ to perform statistical tests and generate publication-quality graphs. For example, I’ve used these tools extensively to analyze the statistical significance of differences in protein expression levels between different treatment groups. I always ensure that appropriate statistical methods are applied and that results are reported accurately and clearly, including any limitations of the analysis.

Troubleshooting a malfunctioning microscope component requires a systematic approach. My strategy starts with a careful examination of the symptoms, including identifying the specific component that’s malfunctioning and the nature of the problem. This might involve checking error messages, visual inspection for any physical damage, and testing the functionality of individual components.

Following a visual inspection, I would systematically eliminate potential causes by testing the microscope’s various systems. For example, if the issue involves image blurriness, I would check the objective lens, condenser alignment, and the focusing mechanism. If the problem involves illumination, I would examine the light source, filters, and pathways. My troubleshooting process also involves checking for loose connections, ensuring proper voltage supply, and considering environmental factors that could affect performance. If the problem persists after these initial checks, I’d consult the microscope’s manual, seek assistance from colleagues with experience, or contact the manufacturer’s technical support for more advanced diagnostics and repairs.

My experience encompasses a wide range of microscopy samples, spanning both biological and material science applications. In the biological domain, I’ve worked with diverse samples including cultured cells (mammalian, bacterial, yeast), tissues (fixed and live), and organisms (whole mounts, sections). This includes experience with both brightfield and fluorescence microscopy applications, requiring different sample preparation techniques for optimal results. For instance, preparing tissue samples often requires fixation, sectioning, and staining protocols optimized for the specific tissue and imaging method.

My experience in material science includes working with various materials including polymers, metals, semiconductors, and nanomaterials. This often requires specialized sample preparation techniques, such as polishing or etching, to achieve appropriate surface finish for high-resolution imaging. The microscopy techniques used in these applications often differ from those used in biological imaging, and I am proficient in selecting appropriate techniques like SEM (Scanning Electron Microscopy) or TEM (Transmission Electron Microscopy) depending on the material properties and information required.

I have extensive experience with various microscopy platforms from leading manufacturers. My experience includes working with inverted and upright microscopes from Nikon (e.g., Eclipse Ti2, A1R confocal), Zeiss (e.g., Axio Observer, LSM 880 confocal), Leica (e.g., DMi8, SP8 confocal), and Olympus (e.g., IX83, FV3000 confocal). This experience allows me to effectively utilize the unique features and functionalities of each platform to optimize experiments for specific applications. For example, I am familiar with the advanced features offered by Nikon’s NIS-Elements software for high-resolution imaging and multi-dimensional data acquisition and analysis, or the automated imaging capabilities of the Zeiss Axio Observer.

Furthermore, I’m familiar with other specialized microscopy platforms such as scanning electron microscopes (SEMs) and transmission electron microscopes (TEMs). My knowledge extends to different types of detectors, light sources, and imaging modes, enabling me to choose the optimal setup for each experiment. This broad platform experience allows me to quickly adapt to new instruments and software, making me a highly versatile microscopist.