Imaging Biological Complexity: A Multimodal Microscopy Approach Using Brightfield, Phase Contrast, and Fluorescence

Abstract

The objective of this lab was to compare brightfield, phase contrast, and fluorescence microscopy in imaging biological samples, assessing their respective advantages and limitations in contrast generation, resolution, and visualization. Key microscopy principles explored included light attenuation in brightfield microscopy, phase shift detection in phase contrast microscopy, and fluorophore excitation and emission in fluorescence microscopy.

The samples examined included histology slides, GFP+ and RFP+ cells, Convallaria rhizome, and 0.5-micron fluorescent beads. Brightfield microscopy provided high-contrast images of stained histology slides but was ineffective for unstained samples. Phase contrast microscopy improved visualization of unstained biological specimens, such as live GFP+ and RFP+ cells, by enhancing refractive index differences. Fluorescence microscopy allowed targeted imaging of cellular structures, using DAPI (nuclear stain), GFP (protein fluorescence), and TRITC (cell wall or vascular marker). Key findings demonstrated that overlaying fluorescence channels provided insight into cellular complexity, but individual channels were necessary to fully distinguish structural components, as seen in Figure 7f, where green fluorescence blocked red signals. Additionally, bead imaging at 40x magnification revealed unexpected variations in calculated bead sizes, likely due to focus-related limitations.

The results highlight the trade-offs between resolution, contrast, and specificity in each microscopy technique. Brightfield microscopy is effective for stained specimens but lacks contrast for unstained samples. Phase contrast enhances visualization of transparent samples but introduces halo artifacts. Fluorescence microscopy enables high-contrast imaging of specific cellular structures but requires careful staining and can suffer from photobleaching. These findings underscore the importance of selecting the appropriate imaging method based on the sample type and experimental objectives.

Introduction

The use of lenses has been in place for centuries – from corrective eyewear to magnifying glasses – to enable us to enhance our vision and see the smaller details. The first use of a simple, single-lens microscope is often credited to Anthony van Leeuwenhoek (1632-1723), leading him to the discovery of tiny protists and bacteria1. That first microscope eventually developed into the standard brightfield microscope we know today, and over the centuries has been instrumental in revealing the structure of cells, the structural bases for the transmission of inherited characteristics, and the microscopic basis of infectious diseases. Brightfield microscopy has also been integral to discoveries in physics and chemistry2. Today, microscopy of sputum smears is the primary method for diagnosing pulmonary tuberculosis in low- and middle-income countries due to its fast and inexpensive nature3. A microscope is also vital in the authentication of medicinal plants. The accurate identification of plant species, variety, and plant part is essential prior to their use in research or treatment4.   

Brightfield, phase contrast, and fluorescence are some of the most common label-free contrast modes used in optical microscopy. In brightfield, or transillumination microscopy, contrast is caused by the difference in light intensity between the sample and the background. Sample illumination is white light transmitted from below and observed from above, and dense areas of the sample attenuates the light and causes visible contrast5. In phase contrast microscopy, the fact that light passing through a specimen travels slower than an undisturbed light beam is used to create contrast. The microscope can detect the phase shifts of colored light relative to each other, and convert this phase shift into something we can see5. In fluorescence microscopy, a sample is illuminated with light of a specific wavelength, which is absorbed by the fluorophores in the sample, causing them to emit light of longer wavelengths, or that of a different color than the absorbed light6. 

The experiments in this report used brightfield, fluorescence, and phase contrast microscopy to look at various sample types using 4x, 20x, and 40x magnification. Histology slides were observed using brightfield at each of the magnifications, and at 20x magnification using phase contrast. Fluorescence imaging was done on DAPI-stained GFP+ and RFP+ cells, 0.5 micron Invitrogen YG fluorescent beads, and convallaria rhizome. Images were taken of various slides and magnifications throughout the lab, and are analyzed below.

This report begins with an overview of the operation of a standard microscope, followed by the imaging and analytic methods used in this lab. Results and associated quantitative and qualitative analyses are presented. Key findings, limitations, and advantages of this lab are summarized in the conclusion. 

Background

Microscopy plays a crucial role in biological and biomedical research, enabling the visualization of microscopic structures with high precision. A fundamental aspect of microscopy is resolution, which determines the ability to distinguish two closely spaced points as separate entities. Resolution is classified into lateral resolution (in the x-y plane) and axial resolution (in the z-plane), both of which are influenced by the numerical aperture (NA) of the objective lens and the wavelength of light used. According to Abbe’s diffraction limit, the smallest resolvable feature size, d, is given by d= λ /2N A​. Axial resolution is typically lower than lateral resolution but can be enhanced using confocal microscopy techniques in fluorescence imaging.

Another critical parameter in microscopy is contrast, which allows differentiation between structures within a sample. Contrast mechanisms vary depending on the imaging technique used. In brightfield microscopy, contrast arises from differences in light absorption by the sample. Phase contrast microscopy utilizes phase shifts due to varying refractive indices to create contrast, making it particularly useful for imaging unstained biological samples. In fluorescence microscopy, contrast is generated by the emission of light from fluorophores excited at specific wavelengths, allowing selective visualization of target structures.

The focal length of a lens also plays a key role in image formation. Defined as the distance between the lens and the point where parallel rays of light converge, focal length directly impacts magnification, with shorter focal lengths providing higher magnifications. In this study, objectives of 4x, 20x, and 40x magnification were used to capture images of histological samples and fluorescently labeled cells. The microscope utilized was an inverted microscope, where the light source and objectives were positioned below the stage, and the detector was placed above. This configuration is particularly advantageous for imaging live cell cultures and thick specimens, as it allows for easy manipulation of samples and reduces interference from coverslips.

Brightfield microscopy, one of the most commonly used imaging techniques, relies on light transmission through the sample, with contrast generated by differences in light absorption. Histology slides were imaged at 4x, 20x, and 40x magnifications using brightfield microscopy to observe cellular structures. To ensure uniform illumination and optimized image quality, Köhler illumination was employed, which aligns the light source and specimen to reduce glare and enhance contrast. Conjugate image planes in this setup included the field diaphragm, condenser, sample, and the image plane of the detector.

Microscopy contrast mechanisms can be classified into absorption-based, phase-based, and fluorescence contrast. Brightfield microscopy utilizes absorption-based contrast, where denser structures appear darker due to greater light attenuation. Phase contrast microscopy generates contrast by detecting phase shifts in light as it passes through samples with varying refractive indices, enhancing visibility of transparent structures. In fluorescence microscopy, contrast arises from fluorophores that absorb specific excitation wavelengths and emit light at longer wavelengths, allowing targeted structures to be visualized with high specificity. 

Phase contrast microscopy was particularly useful in this study for imaging unstained histology slides and GFP+ cells at 20x magnification. This technique relies on specialized components, including a phase plate, which modulates the phase of direct light to enhance contrast, an annular diaphragm that selectively filters light, and an objective with phase rings to match the phase shift induced by the sample.

Fluorescence microscopy is an advanced imaging technique that exploits the properties of fluorophores to generate high-contrast images. In this approach, a high-intensity excitation light source (such as a laser or LED) excites fluorophores within the sample, which subsequently emit light at longer wavelengths. A dichroic mirror selectively reflects excitation light while allowing emitted fluorescence to pass through, and an emission filter ensures only specific wavelengths are detected, reducing background noise.

The fluorescence imaging in this study utilized various filter cubes, each designed to detect specific fluorophores. GFP (green fluorescence protein) excitation occurred at 470 nm, with an emission peak at 525 nm, while DAPI (a DNA-binding stain) was excited at 360 nm and emitted at 460 nm, highlighting cell nuclei. TRITC (tetramethylrhodamine isothiocyanate) was excited at 620 nm and emitted at 700 nm, likely labeling cell walls or vascular components. Fluorescence imaging was conducted on DAPI-stained GFP+ and RFP+ cells, 0.5-micron Invitrogen YG fluorescent beads, and Convallaria rhizome samples at both 4x and 40x magnifications, allowing visualization of specific cellular and structural components.

Overall, the combination of brightfield, phase contrast, and fluorescence microscopy provided a comprehensive approach for imaging biological samples, each technique offering unique advantages based on the sample type and imaging requirements.

Methods

Images were acquired using an inverted fluorescence phase contrast microscope BZ-X810 (KEYENCE, Itasca, Illinois) , which uses a BZ Series infinite optical system. The microscope is configured for Brightfield, Fluorescence (wide-field/sectioning), Phase contrast (PhL, Ph1, Ph2), Oblique illumination. The microscope was equipped with 4X, 20X, and 40X objectives,  a condenser, both trans- and epi- illumination, and filter cubes for DAPI (blue), Texas Red, and GFP (green) fluorescence detection. The microscope has an objective magnification range of 2X to 100X, a working distance range of 0.13 mm to 14.5 mm, and a numerical aperture spanning 0.1 to 1.45. 

We used BZ–X Viewer, which facilitates objective selection, auto-focusing, image acquisition and exposure control.All images were stored in the Biophotonics2025 directory, categorized by magnification and modality, and later analyzed using the Fiji/ImageJ image processing package

Four samples were imaged for this lab: histology slide for brightfield and phase contrast imaging; GFP+ and RFP+ cells stained with DAPI for fluorescence imaging; convallaria rhizome stained for multi-channel fluorescence imaging; 0.5 µm fluorescent beads for resolution and z-stack analysis. All samples were handled with gloves, and at the beginning of the lab, the microscope stage was checked and confirmed to be properly prepared for holding a slide. The laser was powered on using a designated button on the backside of the control box under specific supervision. 

For brightfield imaging, the histology (H&E) stained slide was placed facing downward, as trans-illumination microscopy requires the sample and coverslip to be oriented toward the objective. We captured four images at 4X, 20X, and 40X magnifications, along with a stitched image taken at 40X. Before closing the microscope, we noted the tissue’s location to facilitate easier image acquisition and processing. The BZX–Viewer application was set to ‘Single–Color’ mode, and channel 1 (Brightfield) was selected. We ensured live imaging before proceeding with image capture, and auto–adjusted focus and exposure at each magnification level before manual fine-tuning. Using the ‘Measure’ tab, we added a scale bar to each acquired image before storing the data. To capture a larger area at 40X magnification, a 3x3 composite image centered on the initial field of view was acquired using the XY stitching function. Exposure times were optimized according to the objective lens used: 1/350 seconds for the first objective (4X), 1/120 seconds for the second objective (20X), and 1/30 seconds for the third objective (40X). This is consistent with the physics of the microscopy due to changes in light intensity and a field of view at higher magnifications. As magnification increases from lower to higher (4X to 40X), the field of view decreases, requiring longer exposure times to capture a narrower area with the same ambient light.

To enhance the visualization of unstained cellular structures, we switched from brightfield to phase contrast imaging. We began the process with a histology slide stained with hematoxylin and eosin at 20X magnification, which enabled us to transition to cell imaging. In total, we acquired three images: one in brightfield color mode, one in brightfield monochromatic mode, and one in phase contrast. When switching to phase contrast in the BZX–Viewer application, we selected the ‘Phase Contrast’ setting while maintaining monochromatic imaging. For brightfield (BF) imaging at 20X, the exposure time was recorded as 1/175 seconds, whereas for phase contrast (PC) imaging at the same magnification, it was 1/150 seconds. The slight difference in light exposure can be attributed to the nature of the technique which requires more light to achieve optimal contrast. Phase contrast microscopy relies on differences in refractive index within the sample, which may necessitate slightly longer exposures to capture sufficient contrast information. 

Seven multi-colored fluorescence images were acquired using the ‘Multi-Color’ imaging mode, which was activated with four fluorescence filter channels (CH1–CH4). The fluorescence filter channels were configured as follows: CH1 – brightfield/phase contrast, CH2 – DAPI (blue range), CH3 – Texas Red, and CH4 – GFP (green). To capture images with multiple fluorophore channels simultaneously, we enabled the ‘Multi-Color’ mode in the BZX – Viewer application. To prevent channel overlays during image acquisition, we ensured that the ‘Overlay’ button, located to the right of the channels, remained grey rather than blue.

The samples imaged included GFP-expressing cells, RFP-expressing cells, fluorescent beads (0.5 μm), and Convallaria rhizome samples. Cells were initially located using brightfield imaging at 4X and 10X magnification before switching to fluorescence channels for detailed visualization. Once the cells were located, we switched to 40X magnification and visually inspected to ensure that the cells were not too close to the objective. To facilitate focusing on the cells, we recorded the focus depth (z-depth) at each objective magnification: 4,119 μm for 4X, 3,939 μm for 10X, and 4,112 μm for 40X. We carefully accounted for the exposure time when focusing on cells, as longer exposure times slow down the camera’s response. To protect the cells from photobleaching caused by excessive light and fluorescence exposure, we enabled the ‘Low Photobleach’ setting.

After imaging live cells, we captured high-resolution images of fluorescent beads (0.5 μm) at 20X and 40X magnification. A z-stack image acquisition was performed with a step size (pitch) of 0.2 μm. For bead imaging at 20X, the upper limit was recorded at 3963.4 μm, the lower limit at 3941.8 μm, and the exposure time was 31 seconds. A total of 140 images were captured. For bead imaging at 40X, the focus depth was noted at 3,990.4 μm, with an upper limit of 4003.0 μm and a lower limit of 3973.3 μm. A total of 150 images were acquired, though the exact exposure time was not recorded. To compare high-resolution images with high-sensitivity mode, we increased the exposure by 30% to intentionally induce over-saturation and used the ‘Capture’ button instead of ‘Start Capture’ for bead imaging. Then, we acquired an additional image by switching from ‘High–Resolution’ to ‘High–Sensitivity’ mode. 

For the final stage of this lab, we performed multi-color fluorescence imaging of Convallaria rhizome at 4X and 40X magnification, along with phase contrast (PC) and brightfield (BF) imaging. Notably, we deviated from the lab instructions by using 20X magnification instead of 40X for the stitched image in phase contrast mode. This adjustment was made because we found that 20X magnification provided better structural contrast and was easier to assess.

Results and Analysis

The brightfield histology images captured at 4x (Figure 1a), 20x (Figure 1b), and 40x stitched magnifications (Figure 1c) illustrate how resolution, contrast, depth of field, and working distance change with increasing numerical aperture (NA) and magnification. The primary source of contrast in brightfield microscopy is absorption-based, where different tissue components absorb and scatter light to varying degrees. The H&E staining enhances contrast by selectively binding to cellular structures; hematoxylin stains nuclei dark blue/purple due to its affinity for nucleic acids, while eosin stains cytoplasmic components pink. At lower magnifications (Figure 1a, 4x), the contrast is less pronounced because the larger field of view encompasses both stained and unstained regions. As magnification increases to 20x (Figure 1b) and 40x (Figure 1c), fine cellular structures become more distinct, improving contrast due to enhanced resolution.

The depth of field (DOF), working distance (WD), and exposure time are all affected by magnification and numerical aperture. The DOF decreases as magnification increases, meaning that at 4x (Figure 1a), a larger portion of the sample remains in focus, while at 20x (Figure 1b) and 40x (Figure 1c), precise focusing is required to keep cellular details sharp. Similarly, the working distance decreases with increasing magnification. At 4x, there is ample clearance between the objective and the slide, making it easier to navigate the sample. At 40x (Figure 1c), the working distance is significantly reduced, requiring fine adjustments to avoid losing focus. Exposure time also increases at higher magnifications, as the NA of the objective lens increases, capturing more light but reducing the amount of light reaching the detector. Consequently, the 40x stitched image (Figure 1c) required longer exposure times to compensate for reduced light transmission.

There are clear advantages and limitations to different magnifications. Lower magnifications (4x, Figure 1a) offer a larger field of view and better depth of field, making them useful for scanning and locating areas of interest, but they lack the resolution to observe cellular details. Mid-range magnification (20x, Figure 1b) provides a balance between field of view and resolution, making it ideal for analyzing histological structures while still maintaining a moderate depth of field. Higher magnifications (40x stitched, Figure 1c) allow for detailed visualization of cellular components, but come with a small field of view and shallow depth of field, requiring techniques like stitching to examine larger areas. These trade-offs highlight the importance of choosing an appropriate magnification based on the specific needs of the analysis, balancing resolution, contrast, and field of view accordingly.

For the histology slide at 20x magnification, the brightfield color image (Figure 2a) reveals structural details with natural H&E staining, where hematoxylin highlights the nuclei in blue/purple and eosin stains the cytoplasm pink. This contrast is achieved through absorption-based light attenuation, where stained regions absorb more light than the surrounding tissue. In the brightfield monochromatic image (Figure 2b), color information is removed, leaving only grayscale intensity differences. This enhances contrast between stained and unstained regions by emphasizing variations in light absorption, making subtle intensity shifts more noticeable. The phase contrast image (Figure 2c), however, generates contrast based on differences in refractive index rather than absorption. This allows for better visualization of transparent structures that may not absorb stain well, making it particularly useful for imaging unstained biological specimens. In the PC image, cellular boundaries and organelles appear more defined due to the interference of phase-shifted light waves, enhancing edge contrast.

For the GFP+ cells at 40x magnification, the brightfield color image (Figure 2d) shows some cellular details, but the contrast is relatively low because fluorescence-based signals are not fully optimized under standard transillumination. The brightfield monochromatic image (Figure 2e) improves contrast slightly by eliminating color-based distractions, but still relies primarily on variations in intensity. The phase contrast image (Figure 2f), however, provides significantly improved visibility of unstained cellular structures, as phase contrast microscopy enhances optical path differences in the sample. This results in clearer visualization of cell membranes, internal organelles, and fine structural details, making PC particularly useful for live-cell imaging or unstained biological specimens.

Key differences in image acquisition settings include exposure adjustments for phase contrast to optimize halo effects and fine details, whereas brightfield imaging relies more on illumination intensity and color balance for optimal visibility. The comparison between these imaging techniques demonstrates that brightfield microscopy is ideal for stained samples, whereas phase contrast excels at imaging unstained or weakly absorbing biological specimens by enhancing phase differences rather than absorption. These findings highlight the strengths and limitations of each modality, reinforcing the importance of selecting the appropriate imaging technique based on the sample type and research objective.

The fluorescence images of Convallaria (lily of the valley rhizome) at 4x magnification highlight the differences in contrast mechanisms between brightfield (BF), phase contrast (PC), and fluorescence imaging. The BF image (Figure 3a) provides an overall structural view based on absorption and scattering of light, but lacks specificity in differentiating cellular components. The PC image (Figure 3b) improves visibility by enhancing refractive index differences, making finer structural details more apparent. However, both methods rely on general optical properties rather than specific molecular labeling.

In contrast, fluorescence microscopy provides selective contrast based on fluorophore binding. The DAPI-stained image (Figure 3c) reveals cell nuclei by binding to DNA, emitting blue fluorescence that distinctly highlights nuclear regions. The GFP channel (Figure 3d) shows green fluorescence, likely originating from chlorophyll autofluorescence in the plant tissue or a genetically expressed fluorescent marker. The TRITC (CY5) channel (Figure 3e) produces red fluorescence, potentially highlighting cell wall structures or vascular components within the rhizome. The stitched fluorescence image at 40x magnification (Figure 3f) offers a high-resolution view of the sample, allowing for detailed visualization of fluorescently labeled structures at a finer scale.

Fluorescence imaging provides significantly improved contrast over BF and PC for specific cellular components by utilizing wavelength-dependent excitation and emission rather than relying on variations in optical density or refractive index. This makes it particularly useful for identifying nuclei, proteins, and structural components that are otherwise indistinguishable in conventional microscopy.

Figure 4 shows the lateral full-width-half-maximum (FWHM) intensity profiles of five beads in the selected microscope image. Each bead was measured by drawing a line across the bead in ImageJ, and then plotting an intensity profile along the line. A Gaussian fit function was then used to calculate the FWHM value. Five beads were measured (Table 1) and averaged. A similar process was repeated for calculating the axial FWHM, except a z-stack image of each bead was used to calculate the resolution instead of a single-layer image. The beads were calculated to have an average lateral FWHM diameter of 170.5 nm with a standard deviation of 7.6 nm, and an average axial FWHM diameter of 2729 nm. Both the lateral and axial diameter measurements are very different from the expected 500 nm bead size. 

Using a wavelength () of 500nm for green light and a numerical aperture of 0.6, lateral resolution was calculated to be 416nm and axial resolution to be 2319nm. The measured axial resolution was slightly more than the calculated axial resolution of the microscope. This high resolution could indicate that the image was not fully in focus and thus appeared slightly blurry. To fix this, confocal techniques could be used to eliminate any out-of-focus light. The measured lateral resolution was significantly lower than the calculated lateral resolution of the microscope. This difference could be due to an error in calculations, or if the incorrect pixel size was used during calculation. The resolution could be improved by adjusting the numerical aperture if this experiment was performed again.  

Figure 5 demonstrates that increasing magnification from 20X to 40X enhances the resolution and bead visibility. Figure 5c demonstrates that oversaturation drastically increases fluorescence, but at the cost of detail loss. Finally, the high-sensitivity panel (Figure 5d)improves fluorescence detection without significant loss or oversaturation. 

As shown in Figure 6a, the GFP+ cells provided a poor source of contrast in the brightfield microscopy. The cells are almost indistinguishable from the background, and adjusting the brightness on the microscope did not give better resolution. This suggests that the cells were not dense enough to create detectable lightwave attenuation and contrast. The cells are easily identified through phase contrast microscopy (Figure 6b), however. The light emitted by the microscope passes through the cells, and the wavelength slows enough to be translated into the contrast we can see. In this method, even the light passing through the bubbles in the slide are clearer and easy to identify, whereas brightfield microscopy is still not enough to detect them. Faint shading can be seen within the cells, likely due to components of the cells causing the light passing through them to shift. Fluorescence microscopy was used to observe the cells with the DAPI (Figure 6c), TRITC (FIgure 6d), and GFP (Figure 6e) channels. There was a high amount of GFP detected in the sample, which is expected due to the cells being designed as producers of green fluorescent protein. There is also a small amount of blue detected in the cells, while no red is present. When the brightfield and fluorescent channels are overlaid (Figure 6f), the GFP is the most obvious layer, with faint areas of blue appearing towards the center of the cell.

As shown in Figure 7a, the RFP+ cells provided a slightly better source of contrast in the brightfield microscopy than the GFP+ cells. A viewer may have some difficulty picking out the shades of gray, but the cells are a distinguishably darker shade with a lighter halo surrounding them. It is even possible to detect slightly darker patches within the cells, where denser areas such as organelles may be located. The phase contrast image (Figure 7b) makes these differences even more visible. The cells have clearly-defined edges due to the light haloes, and there are patches of lighter and darker areas within the cells. Cells that were somewhat hard to detect in the brightfield image are now clearly displayed. In this cell slide, fluorescence microscopy was able to detect DAPI (Figure 7c), TRITC (FIgure 7d), and GFP (Figure 7e) fluorophores within the samples. The GFP is the most abundant, but there is a greater presence of blue fluorophores than in the GFP+ cells, as well as a significant amount of red. When the phase contrast and fluorescent channels are overlaid (Figure 7f), the edges of the cells are more clearly defined, and the shading from the overlay gives them a sense of texture. The green is not as prominent as in the GFP+ cells (Figure 6f), and there are stronger areas of blue. There are a few cells that are completely red or shades of reddish-purple. In the RPF+ cells, the red fluorophores are present, whereas they are completely absent in the GFP+ cells. In the Figure 7f overlay, the red color is present, but it is not as strong as the green, so it doesn’t show as well. It’s only when the layers are separated that the viewer can detect how much red is actually present.

Conclusion

In conclusion, this laboratory practice provided us with knowledge on three fundamental laboratory techniques: brightfield, phase contrast, and fluorescent microscopy, highlighting their respective strengths and weaknesses. Simple microscopy is fundamental in quickly characterizing and identifying cells for use in biomedicine, and is available in both low-income and high-income communities, making it particularly appealing. Key findings of this lab demonstrated that brightfield microscopy is ideal for stained specimens, phase contrast enhances visualization of transparent samples, and fluorescence microscopy provides targeted imaging of specific structures. However, brightfield lacked specificity in unstained samples, phase contrast introduced halo artifacts, and fluorescence imaging required specialized staining and photobleaching precautions.

Overall, the lab highlighted the trade-offs in resolution, contrast, and specificity across different microscopy techniques, reinforcing the importance of selecting the appropriate imaging method based on the sample type and research objectives. The findings in this report were possible due to the advanced technology available through cameras, filter cubes, and other components of the modern microscope. When the discovery of tiny protists and bacteria was first made by Anthony van Leeuwenhoek, he likely never envisioned the further advancements in biology, physics, and chemistry that were not possible without that very first single-lens microscope. 

Author Contributions: KS, JRS, and MZ equally contributed to the conceptualization, methodology, investigation, and drafting of the manuscript. JRS and MZ led the revision and editing process, while KS coordinated the final submission. All authors reviewed and approved the final version of the report. MS provided guidance, supervision, and project oversight.

Funding: This work was supported by the Tufts University School of Engineering and made possible through the use of facilities at the Tufts Advanced Microscopic Imaging Center (TAMIC). We thank both institutions for providing access to microscopy equipment and resources as part of the BME-0156: Biophotonics Laboratory course.

Acknowledgments: We sincerely thank Maria Savvidou, Ph.D., Postdoctoral Researcher in the Department of Biomedical Engineering at Tufts University, for her guidance, mentorship, and support throughout the project, particularly in advising and assisting with lab data generation. We also extend our gratitude to Audrey Dutcher, E’25, for her helpful guidance and technical support with microscopy during the laboratory session.

Conflicts of Interest: The authors declare no conflict of interest.

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