# Image Analysis in LabCanvas: a complete guide to every tool and setting, with examples

> Every tool in LabCanvas Image Analysis, tab by tab, with its settings and a real example: image check, display and scale, stacks and Z projection, background subtraction, filters, thresholds and binary operations, ROIs and Set measurements, Analyze Particles, a pixel classifier, spots, cell counter, western blot densitometry, colocalisation, colour deconvolution and eleven guided assays.

Published 2026-11-06 by LabCanvas team. https://www.labcanvas.com/blog/image-analysis-complete-guide

![LabCanvas Image Analysis measuring nuclear translocation: forty numbered cells with their nuclei and cytoplasm rings outlined, and the assay panel with the cell count and the nuclear/cytoplasmic ratio](https://www.labcanvas.com/analysis-examples/translocation.webp)

**LabCanvas Image Analysis** measures microscope images, western blots and gels in the browser, on the raw pixel values. If you have measured images on the desktop before, you will recognise almost everything: an ROI list with **Measure** and **Set measurements**, **Analyze Particles**, rolling-ball **background subtraction**, sixteen automatic **threshold** methods, **Z Project**, **Orthogonal Views**, **Find Maxima**, a **Cell Counter**, gel lanes and a calibration bar. This guide goes through every tab, tool and setting, with a real example of each: every picture below is the app itself, run on one of its built-in sample images, with the numbers it measured in the caption.

## Nanomaterials under the electron microscope

Most of what a nanomaterials lab measures on TEM and SEM images is particle, pore, grain or flake size, and lattice spacing. These are the tools for each, run here on electron micrographs made for the purpose (with a scale in nm, as the microscope writes it):

- **Nanoparticle size distributions from TEM**: **Count** › **Count particles** on the dark particles, with watershed to split touching pairs, a minimum size to drop specks, and **Feret's diameter** in **Set measurements**; send the diameters to Graphs for the histogram and the mean ± SD.
- **Nanorod and nanowire length and width (SEM)**: the same count with **Feret's diameter**: Feret is the length, MinFeret the width, and their ratio the aspect ratio. For branched or curved wires, the **Branching and skeleton** assay gives each branch's length.
- **Lattice spacing from HRTEM**: a line across the fringes in **Measure** › **Profile**, a few pixels wide to average out the noise; count the fringes over the distance for the d-spacing.
- **Pore size and porosity (AAO, porous films)**: count the dark pores with a circularity range for their areas and equivalent diameters; the share of the image on the pore side of the threshold, shown next to the threshold value, is the porosity (the Summary row's %Area counts only the pores that are not cut by the frame).
- **Grain size of thin films**: threshold the grains so that their darker boundaries stay closed (MaxEntropy often suits secondary-electron images; the **Pixel classifier** helps when the contrast is uneven), add watershed only where a boundary is open, then count them for the grain areas and equivalent diameters.
- **Lateral size of 2D flakes**: count the flakes, or train the classifier on flake and substrate, and read the Feret diameters.

![Nanoparticle size from TEM in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/em-tem-np.webp "Nanoparticle size from TEM: 136 gold nanoparticles counted, touching ones split by the watershed: Feret diameter 13.77 ± 1.589 nm mean ± SD.")

![Nanorod length and width SEM in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/em-sem-rods.webp "Nanorod length and width SEM: 36 nanorods those cut by the frame left out: length 496.7 ± 154.6 nm Feret, width 53.47 ± 10.17 nm MinFeret, aspect ratio 11.51 ± 3.885.")

![Lattice spacing from HRTEM in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/em-hrtem.webp "Lattice spacing from HRTEM: Profile across the lattice fringes of a gold nanocrystal: 10 spacings over 2.353 nm, so d = 0.235 nm, the Au 111 spacing.")

![Pore size and porosity AAO in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/em-aao-pores.webp "Pore size and porosity AAO: 160 pores counted those cut by the frame left out: mean area 3,099 nm², an equivalent diameter of 62.8 nm; the pores take up 34.7% of the surface porosity.")

![Grain size of a thin film in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/em-grains.webp "Grain size of a thin film: 118 grains of a perovskite film those cut by the frame left out: mean grain area 39,340 nm², an equivalent diameter of 224 nm.")

![2D flake lateral size in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/em-flakes.webp "2D flake lateral size: 33 MoS₂ flakes those cut by the frame left out: lateral size Feret from 0.3502 to 1.743 µm, 1.043 ± 0.4381 µm on average.")

The rest of this guide goes through every tab, tool and setting.

The app is organised in six tabs on the right (**Image**, **Process**, **Measure**, **Count**, **Blots**, **Assays**), a tool rail on the left, the image in the middle and a results drawer underneath. Sections fold: click a title to fold it, Alt-click to show that section alone.

## Where to find the commands you know

| If you are used to | In LabCanvas Image Analysis |
|---|---|
| ROI Manager, Measure, Set Measurements | **Measure** tab: the ROI list, **Measure**, **Measure all ROIs**, **Set measurements** |
| Analyze Particles | **Count** tab › **Count particles** (fill holes, watershed, size, circularity, exclude on edges) |
| Threshold, Make Binary, Fill Holes, Watershed, Skeletonize, Distance Map, Ultimate Points | **Process** › **Binary**; **Count** › **Count particles** |
| Subtract Background (rolling ball, sliding paraboloid) | **Process** › **Background and filters** › Background |
| Gaussian Blur, Median, Mean, Minimum, Maximum, Unsharp Mask | **Process** › **Background and filters** › Filter |
| Enhance Local Contrast (CLAHE), Remove Outliers, Despeckle, FFT Bandpass | **Process** › **Enhance** |
| Image Calculator, Ratio images | **Process** › **Calculator** |
| Colour Deconvolution, Colour Threshold | **Process** › **Colour deconvolution**, **Colour threshold** |
| Drift and channel registration | **Process** › **Align** |
| Brightness/Contrast, Lookup Tables, Channels Tool, Calibration Bar | **Image** › **Display** |
| Set Scale, Scale Bar | **Image** › **Scale** |
| Z Project, Orthogonal Views, Reslice, Make Substack, Reduce, Animation | **Image** › **Stack and image** |
| Plot Profile, Plot Z-axis Profile | **Measure** › **Profile** |
| Colocalisation (Pearson, Manders, Costes) | **Measure** › **Colocalisation** |
| Enlarge, Make Band, Combine/AND/XOR, Convex Hull, Fit Ellipse, Specify | **Measure** › **ROI tools** |
| Find Maxima | **Count** › **Spots** |
| Cell Counter | **Count** › **Cell counter** |
| Trainable segmentation | **Count** › **Pixel classifier** |
| Batch processing | **Count** › **Batch**, and **Run on many images** in every assay |
| Gel lanes and peaks | **Blots** tab |
| Tracking, kymographs, skeleton analysis, Sholl, angiogenesis networks | **Assays** tab |

## Opening images

Click **Open an image**, press Ctrl O, drop files on the window or paste. Four samples are on the start screen (a western blot, nuclei, two channels for colocalisation and a 15-slice Z-stack) and every assay has its own.

- **TIFF and BigTIFF**: strips or tiles, uncompressed, LZW, Deflate or PackBits; 8- and 16-bit kept as they are, other depths (signed, 32-bit, float) read as 32-bit float. Every page is read; pages of another size (thumbnails, pyramid levels) are left out with a note.
- **Hyperstacks and OME-TIFF**: channels, slices, frames, pixel size, slice spacing, frame interval and channel names come from the file's description. Otherwise the resolution tags are used, but only in centimetres (an inch DPI is a print size, not a pixel size).
- **PNG, JPEG, WebP, GIF, BMP**: read at 8 bits; a grey picture becomes one channel, a colour picture three. JPEG gets a warning that compression changed the pixel values.
- **ROIs**: .roi files and RoiSet.zip (rectangles, ovals, polygons, freehand, traced, straight, segmented and freehand lines, angles, points, composite shapes, with names, colours and positions) go to the image shown.
- **Projects**: a .lcanalysis file holds every image (as TIFF, raw values and scale), its ROIs and all results tables. The session also saves itself in the browser and comes back after a reload.
- Files in a microscope maker's own format (.czi, .nd2, .lif, .lsm, .oib, .vsi, .ims and others) are not read in the browser: export them as TIFF from the acquisition software (OME-TIFF keeps the scale and the channels).

The **File** menu also has **Save image as TIFF** (raw values, hyperstack layout and scale), **Download PNG** (as shown, with the scale bar), **Add to a figure** and **Add channels to a figure**.

### Drawing and editing ROIs

The tool rail: **Rectangle** (R), **Oval** (O), **Polygon** (G), **Freehand** (F), **Line** (L), **Points** (P), **Wand** (W) and **Pan** (H, or hold Space). Shift draws a square or circle. A polygon closes on its first point, a double-click or Enter. The wand traces the object under the click: the red object of the threshold preview when it is on, otherwise the connected pixels within the tolerance set in Count. Click selects an ROI, Shift-click adds, drag moves, handles resize, Delete removes. The readout under the cursor shows x, y, the calibrated position and the value of every channel. Undo keeps 40 steps (Ctrl Z, Ctrl Shift Z).

## The Image tab

### Image check

Is the image fit to measure? The check runs on opening, folds itself away when all is fine and opens when something needs attention:

- **Clipping**: the share of pixels at the top value or at 0, per channel and inside the selected ROIs; **Show where** marks them in red and blue; **Check every slice** goes through a stack.
- **Pixels**: the bit depth and the number of grey levels; flags a picture taken from a figure, a JPEG, or gaps in the histogram left by earlier processing.
- **Illumination**: how much the background varies across the field (a warning above 1.3×), with a link to correct it.
- **Focus**: a sharpness score, and for stacks a bar per slice with the sharpest one marked and **Go to the sharpest slice**.
- **Scale**, and with several images open, **Open images**: a table of their type, pixel size and size that flags images that cannot be compared fairly.

![Image check in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/check.webp "Image check: All fine: nothing clipped brightest value 3,688, even illumination the background varies by 1.06×, sharpest at slice 8 of 15.")

### Display

Display changes only how the image looks. **Composite**, **One channel** or **Gray**; for each channel an eye to hide it, a lookup table (Grays, Red, Green, Blue, Cyan, Magenta, Yellow, Fire, Ice, Viridis, Inferno, Spectrum, Inverted grays), a histogram with the display ramp, minimum and maximum sliders and fields, **Auto** (saturates 0.35% of the pixels at each end) and **Full**. Sliders move through **Channel**, **Slice** and **Frame**. **Colour scale bar** draws a calibration bar for false-colour images, in a corner you choose, with white or black text and a title such as "Ratio 405/488". The footer reminds you which raw bit depth is measured.

### Scale

Draw a line along a scale bar or a micrometer, type the **Known distance** and the **Unit** (µm, nm, mm, cm, m, Å) and click **Set scale**; or type the pixel size directly. **Scale bar** adds a bar with its **Length**, **Position**, **Thickness**, **Text size**, white or black, and a **Label**.

### Stack and image

- **Player**: Play and Pause through frames or slices at a frame rate you set, with **Loop**.
- **Orthogonal views**: XZ and YZ strips through a point you pick; **XZ as image**, **YZ as image** and **Reslice along** a selected line.
- **Z projection**: **Maximum**, **Average**, **Sum**, **Standard deviation**, **Minimum** or **Median**, over the slices you choose, as a new image.
- **Substack and reduce**: a substack of the slices or frames you choose, every n-th, and **Reduce** to keep every n-th plane.
- **Channel as image**, **Crop to rectangle**, and an info list (size, values, planes, pixel size, slice spacing, frame interval, source).

![Z-stack projection in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/stack.webp "Z-stack projection: Maximum intensity projection of slices 1–15 of the bead Z-stack: the beads from every depth sharp in one 256 × 256 image.")

## The Process tab

Every Process step can apply to all channels and planes or only to the ones you choose, optionally **As a new image** and as a **32-bit result**, with progress and Cancel on long stacks. Each step is undoable.

### Background and filters

- **Background**: **Rolling ball radius** (at least the radius of the largest object that is not background), **Light background**, **Sliding paraboloid**, **Smooth first**, a **Preview** of the background and the result, then **Subtract background** or **Background as image**.
- **Filter**: **Gaussian blur** (sigma), **Median**, **Mean**, **Minimum**, **Maximum** (radius) and **Unsharp mask** (sigma and weight).
- **Bit depth**: convert to 8-, 16- or 32-bit, scaled to the display range or not.

### Enhance

For seeing and segmenting, not for measuring intensities (the panel says so). **Local contrast (CLAHE)** with tile size, clip limit and bins; **Denoise (non-local means)** with strength, patch and search sizes, the noise estimated for you; **Remove outliers**, bright or dark, with radius, threshold and a preview of the pixels that would change; **Despeckle**; and **FFT bandpass** (remove structures larger and smaller than sizes you give, optionally horizontal or vertical stripes).

### Binary

On a grey image: pick a **Threshold** method (Default (IsoData), Otsu, Huang, Intermodes, IsoData, Li, MaxEntropy, Mean, MinError, Minimum, Moments, Percentile, RenyiEntropy, Shanbhag, Triangle, Yen, or a value), bright or dark objects, preview it in red, and **Make binary**. On a mask: **Erode**, **Dilate**, **Open**, **Close** (by steps or by a distance in calibrated units), **Fill holes**, **Outline**, **Skeleton**, **Watershed**, **Distance map** and **Ultimate points** (with their radius in a table).

### Calculator

- **Two images**: Subtract, Add, Multiply, Divide, Average, Difference, Minimum, Maximum, AND, OR and XOR between any two channels of open images of the same size, as a 32-bit result if you like; a single-plane image is applied to every plane of a stack.
- **Ratio**: A ÷ B with background values and a threshold estimated for you (or taken from an ROI); dim pixels become NaN, and the 32-bit ratio image opens with the Fire table and a colour scale bar. The median and mean ratio per plane go to a table.
- **One image**: add, subtract, multiply or divide by a value, Log, Exp, Square root, Square, Reciprocal, Invert, or set the background to NaN.

### Align

**Drift**: line each frame (or slice) up with the first, the previous or the mean, measured on a channel you choose, to whole, 1/10, 1/20 or 1/100 pixel, with sub-pixel shifts, on the whole image or inside a rectangle around stable structures. The aligned stack opens as a new image and the drift of every frame goes to a table and a chart. **Channels** measures and corrects the shift between two channels (chromatic shift).

![Drift correction in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/drift.webp "Drift correction: Drift of every frame measured against the first and corrected: largest drift 2.345 µm, at frame 30.")

### Colour deconvolution and colour threshold

For colour pictures of stained sections. **Colour deconvolution** separates **H-DAB**, **H&E** or **H&E-DAB**, or stains you measure yourself on ROIs, into a 32-bit optical-density image with one channel per stain (and a residual channel). When one stain is DAB, a **DAB positive** block gives the DAB-positive area and the mean DAB optical density, with a threshold set automatically (or by you), per ROI or for the whole image. **Colour threshold** selects pixels by **Hue, saturation, brightness** or **L\*a\*b\*** around colours you pick (Shift-click adds a colour) with a tolerance, then measures the colour area, makes ROIs or a mask.

![Colour deconvolution IHC in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/ihc.webp "Colour deconvolution IHC: Hematoxylin and DAB separated by colour deconvolution: 8.41% of the section DAB positive, mean DAB OD 0.0552 0.483 in the positive pixels.")

## The Measure tab

### Measure and Set measurements

The ROI list shows every ROI with its kind, name and size: click to select, Shift or Ctrl to add, double-click or F2 to rename, arrows to move. **Import…** and **Export RoiSet.zip** move ROI sets in and out. **Measure** measures the selected ROIs (or the whole image), **Measure all ROIs** measures every one, on **This channel** or **All channels**, and on this plane or **Every slice** or **Every frame**. **Set measurements** chooses the columns:

- **Area**, **Mean**, **Min and max** and **Integrated density** (on by default);
- **Standard deviation**, **Median**, **Centroid**, **Centre of mass**, **Perimeter**;
- **Shape descriptors** (circularity, aspect ratio, roundness, solidity), **Fit ellipse** (major, minor, angle) and **Feret's diameter** (Feret, MinFeret, angle and start point).

The table columns are the familiar ones: Area, Mean, StdDev, Min, Max, Median, IntDen, RawIntDen, X, Y, XM, YM, Perim., Circ., AR, Round, Solidity, Major, Minor, Angle, Feret, MinFeret, FeretAngle, FeretX and FeretY. Lines give their length and angle and the values along them; points give their count.

### ROI tools

- **Combine**, **Intersect**, **XOR** and **Subtract** (the first minus the others).
- **Enlarge**, **Shrink** and **Band** (a ring; rings of neighbours are split halfway), by a distance in pixels or calibrated units.
- **Hull**, **Ellipse**, **Circle** (fitted) and **Split** (a composite ROI into its pieces); **Rotate** and **Scale**.
- **Mask image** from the ROIs; **ROIs from a mask image** (objects or label values); and an **Exact position and size** dialog for rectangles and ovals.

### Profile

With a line selected, a live plot of the values along it, one curve per channel, with a **Line width** for a wider average; hovering the plot marks the point on the image. With no line on a stack, the mean of the selected area through slices or through time (a Z-axis or time profile). **Add to results** puts the numbers in a table.

![Line intensity profile in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/profile.webp "Line intensity profile: Green and magenta values along a 28.56 µm line Line 1, 3 px wide, one curve per channel.")

### Colocalisation

Choose two channels, a region (the selected ROI or the whole image), all slices of a Z-stack or one, and the thresholds: **Costes (auto)**, values you set, or none. **Calculate** draws the 2D histogram with its regression line and gives **Pearson's r**, r above the thresholds, **Spearman's ρ**, **Manders M1 and M2**, thresholded **tM1 and tM2**, the overlap coefficient and **Li's ICQ**; **Show colocalised** marks the pixels above both thresholds in yellow. Each run adds a row to the Colocalisation table.

![Colocalisation in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/coloc.webp "Colocalisation: Pearson's r 0.529; above the Costes thresholds tM1 0.219 and tM2 0.214 7,169 colocalised pixels, in yellow.")

## The Count tab

### Count particles

The everyday **Analyze Particles**. Objects come from a **Threshold** (any of the sixteen methods or a manual cut set by dragging on the histogram, bright or dark objects, on a channel you choose, previewed in red) or from the **Classifier** below. Options: **Fill holes**, **Separate touching objects (watershed)**, **Exclude particles on the edges**, **Only inside the selection**, a size range, a circularity range, sizes in pixels or calibrated units, and which image to **Measure** (the thresholded channel, another channel or another open image of the same size). **Count** outlines and numbers every particle and fills a Particles table and a Summary row (Count, Total Area, Average Size, %Area, Mean); **Add as ROIs** sends them to the ROI list; **Find nuclei** does Otsu, fill holes, watershed and a sensible minimum size in one click.

![Counting nuclei in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/nuclei.webp "Counting nuclei: 70 nuclei counted Otsu threshold, watershed to split touching ones: mean area 40.47 µm², 10.2% of the field.")

### Pixel classifier

When a threshold cannot separate what you want, paint a few strokes on each class (Object and Background to start, up to six classes) and a random forest of 100 trees colours the rest, live; paint more where it is wrong. The brush has a size and an eraser (or hold Alt); the view shows the **Result**, the **Uncertain** pixels or your **Strokes**. **Count objects** counts with the Count particles settings, **Make ROIs** makes ROIs, **Probability image** gives one 32-bit channel per class and **Areas** measures the area of each class. **Detail** (fine, medium, coarse) sets the scale of the features it looks at; a classifier can be saved and loaded.

![Pixel classifier in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/classifier.webp "Pixel classifier: Trained on 1,018 painted pixels: 70 objects counted, the Object class covering 12.3% of the image.")

### Spots

**Find Maxima** for puncta, foci and beads: bright or dark spots, a **Prominence** set automatically from the image's noise (or by you), **Smoothing**, **Skip edge spots**, counted in the selected ROIs or over the whole image, with a live preview of every spot. The table gives the count, area and density per ROI, and optionally every spot's position and value; **Add as point ROIs** keeps them.

![Spot detection in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/spots.webp "Spot detection: 150 spots found in the green channel, with the prominence 132.5 set automatically from the image's noise.")

### Cell counter and Batch

The **Cell counter** counts by clicking, in up to 20 named categories (keys 1 to 9 switch between them; a click on a marker removes it), and gives the count, total and percentage per category, with optional marker positions and their values. **Batch** runs the Count particles settings on every open image or on files you choose, opened one at a time: one Summary row per image, and every particle if you want them.

## The Blots tab

Western blot and gel densitometry in three steps. **Band look** (dark or light bands) is detected from the image.

1. **Lanes**: vertical or horizontal; **Find lanes**, **Add lane** or **Use selected** rectangles, the same width for every lane. Choose a **Molecular-weight ladder** lane and type its sizes (for example 250, 150, 100, 75, 50, 37, 25, 20, 15, 10 kDa), with a straight or curved fit, to read every band's size.
2. **Bands**: **Find bands** proposes the band rows across lanes; drag their windows on the image or on the lane plot, name them and mark the loading control. **Align lanes** corrects smiling or shifted lanes before bands are matched.
3. **Quantify**: a **Background** under each band (**Straight line**, **Rolling** or **None**), **Normalise to** a loading-control band or a **Total protein stain** image (Ponceau S, stain-free, with an offset if it is not aligned), and a **Reference lane**. The table gives each band's area, its value relative to the reference lane, the normalised value and the fold change, the lane's total protein and the band's molecular weight; it updates as you change anything. **Lane profile** shows each lane's plot with its band windows.

![Western blot densitometry in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/blot.webp "Western blot densitometry: 8 lanes; the target band normalised to the loading control rises from 1 lane 1 to 8.885-fold lane 8.")

## The Assays tab

Guided measurements for common experiments. Each opens as numbered steps with settings that work in one click, shows its result on the image as you change a setting, offers **Try it on a sample**, and ends in tables you can send to Graphs. **Save as workflow** keeps an assay's settings under a name, listed at the top of the tab; a workflow runs on every open image or on files (**Run on N open**, **Files…**), and the results collect in one table, a row per image or per cell.

### Nuclear vs cytoplasmic signal

For NF-κB, YAP or TFEB translocation (two channels: nuclei and signal). Nuclei are found with a threshold (Otsu by default), blur, a smallest size, fill holes, split touching nuclei and leave out nuclei on the edge; a cytoplasm ring is drawn around each with an **Inset**, **Ring gap** and **Ring width** (rings never overlap a neighbour, and can be kept inside the cell). Background: none, automatic, a value or a region. Per cell: nucleus and ring area, mean and integrated density, the **N/C ratio** and the nuclear fraction, plus a summary.

![Nuclear vs cytoplasmic signal in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/translocation.webp "Nuclear vs cytoplasmic signal: 40 cells, nuclear/cytoplasmic ratio 1.64 ± 1.1, nuclear fraction 0.53 ± 0.21.")

### Spots per cell

For γH2AX, 53BP1 or LC3 foci. The same nuclei step, then spots with optional background removal, blur and a prominence set from the noise; counted in the nucleus or in the whole cell (a distance around the nucleus), with the peak value or a disc mean as intensity, and a number of spots that makes a cell positive. Per cell: spots, mean and total spot intensity, spots per area; and the share of positive cells.

![Spots per cell in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/foci.webp "Spots per cell: 30 cells, 6.63 ± 4.3 spots per cell, 60% of cells with 5 or more.")

### Colony counting

In a **Round dish** (found automatically, leaving out a rim), in **Drawn wells**, or over the whole image; dark-on-light or light-on-dark colonies, a threshold, a smallest colony, fill holes and split touching colonies. Gives the count, colonies per cm² on a calibrated image, mean size and diameter and the area covered, and if you like every colony's size.

![Colony counting in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/colonies.webp "Colony counting: 60 colonies 2.21 per cm², mean size 1.26 mm², 2.79% of the dish covered.")

### Wound healing (scratch)

The open area of a scratch from the image's texture (a window radius and a threshold, the same for every frame), keeping the region that reaches the edge, the largest or all, and filling specks inside it. On a time-lapse, **Measure all frames** gives the open area, % of initial area, % closure and mean width over time, and the closure rates (area and width per hour, with their R²); one image gives its open area and width.

![Wound healing scratch in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/wound.webp "Wound healing scratch: Scratch 75.2% closed by the last frame, closing at 4,000 µm²/h R² 1.000.")

### FRAP recovery

The bleach frame is found automatically; the bleached spot, the reference (whole cell) and the background are found too, or taken from ROIs you draw. The recovery is double normalised, scaled to full scale and fitted with \(I(t) = I_0 + A \cdot (1 - e^{-kt})\): mobile and immobile fraction, half-time t½, k, R², bleach depth and gap ratio.

![FRAP recovery in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/frap.webp "FRAP recovery: Mobile fraction 0.8, recovery half-time 2.77 s, fit R² 0.9999.")

### Calcium traces (ΔF/F₀)

Cells found on a standard-deviation, maximum or mean projection (or your own ROIs), background removed, a baseline F₀ from a low percentile or the first frames, and peaks above a height, a rise and a minimum spacing. Per cell: area, F₀, number of peaks, maximum ΔF/F₀, first peak, frequency and area under the curve, plus every trace in one table and an interactive chart.

![Calcium traces ΔF/F₀ in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/calcium.webp "Calcium traces ΔF/F₀: 20 cells traced, 20 with peaks, 6.1 peaks per minute on average.")

### Track particles and cells

Detect spots or objects in every frame, then link them into tracks with a largest move, missed frames allowed and a shortest track. Per track: duration, path length, net displacement, mean and maximum speed and straightness; the mean squared displacement (with the drift subtracted if you like) is fitted for the diffusion coefficient D. Tracks are drawn in colour on the image.

![Particle and cell tracking in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/tracking.webp "Particle and cell tracking: 40 tracks, speed 0.358 ± 0.022 µm/s, diffusion coefficient D 0.0392 µm²/s.")

### Kymograph

Draw or trace a line along a filament or axon (or let **Find the track** find the brightest one), set its width and **Make kymograph**: distance across, one row per frame. Then draw lines along the streaks: each gives its velocity, and the summary splits them into moving towards the end, towards the start and still.

![Kymograph in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/kymograph.webp "Kymograph: 5 particle streaks traced on the kymograph: velocities from -1.42 to 1 µm/s, speed 0.684 ± 0.55 µm/s mean ± SD.")

### Branching and skeleton

For neurites, vessels, hyphae or filaments: find the structures with a threshold (Li by default), smoothing, smallest object and holes to fill, prune short spurs, and **Analyse**. Gives skeletons, branches, junctions, end points, triple and quadruple points, total, mean and maximum branch length, and every branch with its length, straight-line distance and tortuosity.

![Branching and skeleton in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/branching.webp "Branching and skeleton: 41 branches, 19 junctions and 24 end points; total length 390.6 µm.")

### Sholl analysis

The cell is found as for branching, counted on the cell or on its skeleton, around the cell body (found automatically) or a centre you click, on circles from a start to an end radius in steps you set. Gives the Sholl profile, the maximum number of crossings and its critical radius, the sum and mean, primary branches, the ramification index and the Sholl regression coefficient k with its R².

![Sholl analysis in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/sholl.webp "Sholl analysis: Up to 17 branch crossings, on the circle 30 µm from the cell body; ramification index 2.83.")

### Tube formation

Endothelial networks on matrix, in phase contrast or fluorescence: the tubes are found (with background flattening), short sprouts pruned and meshes smaller than a size left out. Gives total tube length, segments, branches, isolated elements, junctions, meshes and their total and mean area.

![Tube formation in LabCanvas Image Analysis](https://www.labcanvas.com/analysis-examples/tubes.webp "Tube formation: Total tube length 15,190 µm, 123 junctions, 169 segments, 46 meshes.")

## Results, and where they go

Every table sits in the results drawer under the image, one tab per table, with units in the column headers. Click a row to select its ROI and image. Above each table:

- **Summary** adds Mean, SD and n rows; **Copy** puts the table on the clipboard for a spreadsheet; **CSV** saves it; **Clear** empties it.
- **Send to Graphs** makes a Graphs data table with a graph from the column you pick: **By image** (one data set per image, for example control and treated), **All rows**, or **XY** for profiles, time series and other tables with a position column. Sending the same table again updates the same graph.
- **Methods**: each table carries a sentence describing how it was measured (on which values, with which definitions and pixel size), ready to copy into the methods section.

Images go to figures too: **Add image to canvas** places the image as shown, with its scale bar (and its ROIs and outlines if you tick that), in the figure, slide or poster open in another tab; **Each channel and the merge** sends every channel and the merge as separate pictures side by side for a panel layout, single channels in grey if you like.

## Keyboard shortcuts

| Keys | Action |
|---|---|
| R, O, G, F, L, P, W, H | Rectangle, Oval, Polygon, Freehand, Line, Points, Wand, Pan |
| Space (hold) | Pan |
| Ctrl Z, Ctrl Shift Z | Undo, redo |
| Ctrl O, Ctrl S | Open, save the project |
| +, −, 0, 1 | Zoom in, zoom out, fit to window, 100% |
| Delete | Remove the selected ROIs |
| F2 or Enter | Rename the selected ROI or category |
| 1 to 9 | Switch categories in the cell counter |
| Esc | Stop a tool, cancel a polygon, deselect |

## Good practice

- Measure the original file from the instrument, not a picture exported for a figure: JPEG compression and adjustments change pixel values. The image check flags both.
- Keep acquisition settings the same for images you compare, avoid clipped pixels, and measure them with the same settings: saved workflows and Batch make that easy.
- Enhance and denoise to see and to segment; measure intensities on the original image.
- For blots, stay within the linear range of the detection and normalise to a loading control or total protein.

Every measurement here is checked against independent software and against images with known answers, in more than 4,000 automated tests: see [how Image Analysis is validated](https://www.labcanvas.com/image-analysis-validation). Ready to try it? [Open Image Analysis](https://www.labcanvas.com/analysis.html) and click a sample.

## Frequently asked questions

### Will it feel familiar if I already measure images on the desktop?

Yes. The tools carry the names you know: an ROI list with Measure and Set measurements, Analyze Particles with fill holes, watershed, size and circularity, rolling-ball background subtraction, the usual automatic threshold methods (Default IsoData, Otsu, Li, Triangle, Huang, MaxEntropy, Yen and the rest), Z Project, Orthogonal Views, Reslice, Find Maxima, Cell Counter, Plot Profile, gel lanes, colour deconvolution, a calibration bar and a scale bar. ROI sets (.roi, RoiSet.zip) and hyperstack TIFFs open and save in the usual formats.

### Do the display settings change my measurements?

No. Brightness, contrast and lookup tables change only how the image is shown. Every measurement, threshold, assay and blot reads the raw 8-, 16- or 32-bit values, and the image check warns when the data are clipped, compressed or came from a figure rather than the instrument.

### Which threshold methods are there?

Default (IsoData), Otsu, Huang, Intermodes, IsoData, Li, MaxEntropy, Mean, MinError, Minimum, Moments, Percentile, RenyiEntropy, Shanbhag, Triangle and Yen, plus a value you set by hand or by dragging on the histogram.

### Can I run the same measurement on a whole folder of images?

Yes. Counting has a Batch section that runs the Count particles settings on every open image or on files you choose. Every guided assay except the kymograph can be saved as a workflow and run on many images; the files are opened one at a time and the results collect in one table, one row per image or per cell.

### Which files open?

TIFF and BigTIFF (stacks, hyperstacks, 8-, 16- and 32-bit, OME-TIFF), PNG, JPEG, WebP, GIF and BMP, ROI files (.roi, RoiSet.zip) and LabCanvas projects (.lcanalysis). Files in a microscope maker's own format (.czi, .nd2, .lif and others) need to be exported as TIFF first; OME-TIFF keeps the scale and channels.

## Related links

- [Open Image Analysis](https://www.labcanvas.com/analysis.html)
- [Image Analysis at a glance](https://www.labcanvas.com/blog/image-analysis-microscopy-western-blot): The short overview of the app
- [How Image Analysis is validated](https://www.labcanvas.com/image-analysis-validation): More than 4,000 checks against independent software and images with known answers
- [Graphs & statistics guide](https://www.labcanvas.com/blog/graphs-statistics-scientific-plots): Tests and graphs for the numbers you measure
- [The Pixel editor guide](https://www.labcanvas.com/blog/pixel-editor-microscopy-images): Adjust images for the figure, after measuring