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35 protocols using image processing toolbox

1

Estimating Intracellular Diffusion Dynamics

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The whole data analysis workflow was developed and run in Matlab (MathWorks) using the Image Processing Toolbox. After importing the image files as data matrices, the contribution to correlation due to immobile background and slowly moving intracellular structures was removed by applying the moving average filter, as described elsewhere21 (link). The average correlation matrix is calculated according to equation 2 and fitted to equation 15 (Supplementary Information) in order to obtain the iMSD. We use this procedure as an algorithmic way to obtain an estimate of the width of the iMSD distribution for each scan speed. The whole process is repeated for every measured pixel dwell time in order to obtain the complete iMSD. All measurements in which the cell significantly moves or bright vesicles cross the ROI are discarded from analysis to avoid artefacts. The iMSD points shown in the figures were obtained by averaging couples of adjacent points to reduce the noise.
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2

Quantitative Analysis of Neuronal Dendrites

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Segmentation and volume rendering of neurons and neuropils were carried out in AMIRA 6.2 (FEI, Hillsboro, OL). To quantify the dendritic volume, we counted the number of voxels with segmentation data of individual neurons. We excluded soma and axon-like processes for subsequent analysis. The volume distribution of dendrites (Fig. 3D,E) was calculated using a custom-made program written in Matlab and an image processing toolbox (MathWorks, Natick, MA). Maximum intensity projection images were prepared with ImageJ45 . Figures were prepared in Adobe Illustrator CS (Adobe Systems, San Jose, CA). The datasets generated during and/or analysed during the current study are available from the corresponding author on reasonable request.
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3

Automated Conjunctival Microvascular Assessment

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The automated technique for hemodynamic assessment of the conjunctival microvascular network consisted of several image processing steps, as depicted in the flow chart of Fig. 1. Briefly, the automated approach consisted of image registration for correction of eye movement, image segmentation to identify vessels, centerline extraction and bifurcation detection to define centerlines of individual vessel segments, diameter measurement, detection of blood flow, and measurement of axial blood velocity. All image processing and analysis algorithms were developed in Matlab (Release 2014a, MathWorks, Inc., Natick, MA, USA) with image processing toolbox version 9.0. Further detail on the analysis steps is provided below.
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4

Geocorrection Method for Field Data

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To minimize the impact of the hardware issues encountered during the 2017 field campaign, an alternative geocorrection method was developed (Figure 2). The methodology can be divided into the following steps: Databoss alignment, initial geolocation, frame registration, and gridding. This methodology was developed and performed using MATLAB Version 2016a, and the Image Processing Toolbox (The MathWorks, Inc., Natick, MA, USA). The specific code for this process can be found in [38 ].
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5

Automated Quantification of Stromal Nerves

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MxIHC whole slide image (WSI) extraction, registration, and layering was performed in MATLAB with the Image Processing toolbox (The MathWorks, Inc., Natick, MA, USA) (outlined in supplementary material, Figure S1). Registered and extracted WSIs were tiled for ease of processing and were loaded into ilastik 1.3.2 [30 (link)] to segment the stromal component. In brief, subregions of these tiles were flipped and rotated as the training material. Tissue, non-tissue, stromal, and non-stromal masks were generated by training on the hematoxylin and vimentin layers. Using these parameters, tiles were segmented into “glass”, “epithelium”, “stroma”, and “parenchyma” by training with oversight by a pathologist. The “parenchyma” component was expanded to eliminate stroma located in other normal tissue. To identify tissue structures within the images, an automated pipeline was created in CellProfiler [31 (link)]. Within the stromal mask, structures positive for both Gap-43 and S100, measuring >500 pixel2 (equivalent 11 μm2) were counted as nerves, while objects smaller than 500 pixel2 were considered as Gap-43+/S100+ small objects. This method is described in Supplementary materials and methods.
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6

Implant Failure Evaluation Protocol

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Time, number of cycles, axial load and displacement were recorded using the machine transducers with a frequency of 64 Hz. The axial stiffness was calculated from the load–displacement curves of cycles 10 to 19. Therefore, minimal and maximal force, as well as displacement values have been used. For further evaluation, the mean value was used. Every 250 cycles, an x-ray in anterior–posterior view, including a reference sphere for scaling, was taken at the base load of 100 N. X-ray evaluation was performed using a custom-made software routine (Matlab 7.9 R2009b, Image processing Toolbox, The MathWorks GmbH, Ismaning, Germany). The number of cycles until 5°varus collapse compared with the initial x-ray was identified for all specimens and defined as number of cycles to failure [8 (link)]. Specimens were tested until catastrophic implant or construct failure.
After testing normal distribution of the data (Shapiro–Wilk test), the Wilcoxon signed-rank test was carried out to identify differences between study groups regarding axial stiffness and cycles to failure. The software package SPSS 24.0 (IBMN, IBM SPSS Statistics, Version 24.0, New York, US) was used for all statistical evaluations. The level of significance was set to alpha = 0.05.
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7

Viscoelastic Cell Deformation Dynamics

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Obtained sequential images were processed in a custom routine developed in MATLAB R2015a with Image processing toolbox (MathWorks). Using the image in the catching phase, data screening was conducted for the cell with high ellipsoidal eccentricity ( > 0.5) to avoid the analyses of unexpected asymmetric deformation inside the narrow path. Then the cell shape and height in the launching phase was measured based on the binarized images calculated from the original bright field images. The changes in cell orientation after launch was corrected in every image before the height calculation based on the angular degree of the ellipsoidal minor axis of the binarized cell. Least square fittings of the cell height changes for various loading time T = 0 s, 5 s, 15 s, 30 s, 45 s, 60 s, 120 s, 180 s, and 300 s with equation (4) was performed using the mean cell height data after 80 ms (standard saline and that with ATP-depletion assay) and for T = 0 s, 5 s, 15 s, 60 s, 180 s, and 300 s after 100 ms (standard saline dissolving 100 μg/ml S-LPS) when the cells stopped initial elastic responses and entered the viscoelastic regime in the launching phase.
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8

Quantitative Analysis of Fibrin Matrices

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Z Stack images of fibrin matrices were visualized and quantified using multiple feature detection techniques of the Image Processing Toolbox in MATLAB (MathWorks).51 Geometrical tubular elements were identified utilizing the fibermetric function, which employed multiscale second-order local structures (a vesselness measure) of eigenvalues from an Hessian pixilated matrix to detect tubular structures.52 This function was used to identify tubular structures and was also used to filter out areas of dense fibrin fiber overlap. A mean filter function was used to identify densely populated fiber areas within each original z stack slice. These 2 preprocessing techniques (Figure 1) were combined into a single preprocessed image and used in subsequent segmentation and image-processing methods. Specifically, houghpeaks (Hough transform) and houghlines (Hough line) functions were applied to generate Cartesian line segments through the center of tubular structures to detect fibrin fibers.46 (link),52 ,53 (link) Overall, fibrin fiber overlap and fibrin fiber length were calculated after Hough line generation, while fiber matrix porosity and fractal dimension were computed from preprocessed 2-D slices prior to segmentation.
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9

Quantitative Analysis of Collagen Fiber Networks

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Images of Picrosirius Red-stained slides were captured at 40 × and then color deconvoluted, converted to gray scale, binarized, and skeletonized using a novel algorithm run in MATLAB with Image Processing Toolbox (R2018b, MathWorks, Natick, MA)41 . From the skeletonized images, 13 parameters of red collagen fibers were extracted (brightness, number, length, width, persistence, angle, branchpoints, euler number, extent, perimeter, solidity, eccentricity, equivalent diameter), measured using the regionprops command42 (link), and underwent dimensionality reduction to generate 2 dimensional t-Distributed Stochastic Neighbor Embedding (TSNE) plots to visualize collective differences in the collagen fiber network patterns between groups. Geometric shapes were drawn to approximate the distribution of each cluster.
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10

Single-Cell Blood Flow Measurement

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Single-cell blood flow was measured using our recently published approach (Joseph et al., 2019 (link)), with custom code written in MATLAB R2017a (Version 9.2, with Image Processing Toolbox, MathWorks, MA). Source code is available in public repository here: Joseph, 2020
https://github.com/abyjoseph1991/single_cell_blood_flow (copy archived at https://github.com/elifesciences-publications/single_cell_blood_flow_2). AOSLO dataset has been made available at a public repository here: https://doi.org/10.5281/zenodo.2658767.
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