A Simple Method for Quantifying Blastema Growth in Regenerating Planarians

Abstract

Due to their strong regenerative capabilities, freshwater planarians are a well-suited model system for studying the effects of chemicals on stem cell biology and regeneration. After amputation, a planarian will regenerate the missing body parts within 1 to 2 weeks. Because planarians have a distinct head morphology that can be easily identified, head and eye regeneration has been a popular qualitative measure of toxicity. However, qualitative measures can only detect strong defects. Here, we present protocols for quantifying the rate of blastema growth to measure regeneration defects for assessment of chemical toxicity. Following amputation, a regenerative blastema forms at the wound site. Over the course of several days, the blastema grows and subsequently reforms the missing anatomical structures. This growth can be measured by imaging the regenerating planarian. As the blastema tissue is unpigmented, it can be easily distinguished from the remaining pigmented body using standard image analysis techniques. Basic Protocol 1 provides a step-by-step guide for imaging regenerating planarians over several days of regeneration. Basic Protocol 2 describes the necessary steps for the quantification of blastema size using freeware. It is accompanied by video tutorials to facilitate adaptation. Basic Protocol 3 shows how to calculate the growth rate using linear curve fitting in a spreadsheet. The ease of implementation and low cost make this procedure suitable for an undergraduate laboratory teaching setting, in addition to typical research settings. Although we focus on head regeneration in Dugesia japonica, these protocols are adaptable to other wound sites and planarian species.

INTRODUCTION

Freshwater planarians have been a popular model for pharmacological and toxicological studies of chemical effects on regeneration and behavior for decades (reviewed in Best & Morita, 1982; Hagstrom, Cochet-Escartin, & Collins, 2016; see Current Protocols article: Ireland & Collins, 2022; Wu & Li, 2018). Most species have a distinct head morphology with anatomical structures that can be easily identified (eyes and auricles) and manually scored to evaluate possible effects of chemical exposure on regeneration. Following head amputation, a regenerative blastema forms at the wound site. Over the course of several days, the blastema enlarges. Within the blastema, stem cells proliferate and differentiate to regenerate the missing body parts (reviewed in Ivankovic et al., 2019). Eyes and auricles reappear around 5 to 7 days post-amputation. Gross regeneration defects such as missing or extraneous eyes or defects in auricle formation have been a popular readout of chemical systemic or developmental toxicity (as, for example, in Sheehan & Hoegler, 2018). However, more subtle regeneration defects, such as slowed growth or aberrant tissue, are likely to be missed with such a qualitative analysis. This limits the application of using regeneration kinematics as a readout to evaluate chemical safety. To overcome this limitation, we have developed a low-cost and easy-to-implement method to quantify blastema growth. Following amputation on day 1, the regenerating planarian is imaged daily starting on day 4 using a simple light microscope. The acquired images are then processed in free image analysis software (ImageJ; Schindelin et al., 2012). As the blastema tissue is unpigmented, it can be distinguished from the remaining pigmented body using standard image analysis techniques. Because the technique measures the blastema tissue over the course of regeneration starting prior to eye reappearance, it can detect regeneration delays earlier. Thus, blastema growth measurements complement eye-scoring assays.

Basic Protocol 1 is a step-by-step protocol on how to image planarians during regeneration to allow for quantitative analysis. Basic Protocol 2 explains how to use the acquired images to quantify blastema size using ImageJ (Schindelin et al., 2012). Basic Protocol 3 explains how the measured areas from the different days are normalized by the original size of the planarian and used to calculate the blastema growth rate using linear curve fitting. This method could also be used to study the effects of gene knockdown (RNA interference; RNAi) or different environmental conditions on regeneration kinematics or on different wound sites other than head amputation sites. We provide a spreadsheet in the Supporting Information that contains example data and an example workflow and that can be used as a template for the analysis.

STRATEGIC PLANNING

Planarians should be starved for 3 to 5 days before the start of the experiment. At least 10 specimens should be used per treatment condition. Basic Protocol 1 is a multi-day experiment. To be able to analyze blastema growth, several images are taken on different days. It is recommended to amputate the planarians on Friday (day 1) and to image daily on Monday through Thursday (days 4 to 7). Imaging prior to day 4 is not informative, as the regenerated blastema is too small to be imaged robustly using these imaging conditions. This timeline may require adjustment depending on the species or treatment used. It is recommended that images are quality control–checked during imaging to ensure analysis is possible. If necessary, images can be retaken on the same day. It is recommended to take multiple (2 to 3) images of the same worm to ensure images of adequate quality are obtained. Imaging will take about 30 to 60 min per 10 worms, with variation based on experience level. Basic Protocol 2 (image analysis) and Basic Protocol 3 (growth rate analysis) can be conducted separately from imaging. The timeline for these protocols is flexible, but it is best to analyze all images from a single experiment together for consistency.

BASIC PROTOCOL 1 IMAGING PLANARIANS DURING REGENERATION

This protocol describes the steps for imaging planarians over consecutive days of regeneration using a light microscope. On day 1, intact planarians are imaged before they are decapitated to allow for scaling by the original size of the planarian. The anterior pieces are discarded, whereas the regenerating posterior pieces are subsequently imaged daily on days 4 to 7. Emphasis is given to the acquisition of high-quality images using a uniform dark background to allow for quantitative image analysis in Basic Protocol 2. Because planarians are light aversive (photophobic), they will move a lot during imaging. Thus, it is recommended to use a larger imaging dish (e.g., 100-mm petri dish) to be able to keep the planarian in the field of view and away from the boundaries of the container for longer and thus increase the success rate of image acquisition. If chemical exposure is performed, a wash step is included to allow imaging in planarian water and thus minimize exposure to chemicals. Use special care to ensure the planarians are kept in the same order so that the identity of each planarian is known throughout the experiment.

Materials

Planarians of similar size from species of choice (e.g., Schmidtea mediterranea, Dugesia japonica, Girardia dorotocephala; minimum of 10 worms per experimental group)

Planarian water [1× Montjuic Salts (Cebrià & Newmark, 2005), 1× Instant Ocean (IO; 0.5 g/L; Spectrum Brands), or bottled spring water; ≥80 ml will be needed on day 1 for imaging and amputation, 20 ml will be needed each regeneration day for imaging for each experimental group, and additional 20 ml will be needed to wash chemically exposed planarians]

10× stock of desired exposure concentration of chemical (2 ml working solution will be needed for 10 planarians)

100-mm petri dishes (for imaging; four dishes on day 1 and one dish per chemical treatment for days 4 to 7)

Plastic transfer pipets (Samco 691–1S or equivalent; to handle worms; two pipets per chemical treatment)

Light microscope, with camera and data-recording capacity

12-well tissue culture–treated plates (or equivalent containers, for storing planarians when not being imaged and for wash steps; three plates per chemical treatment)

Ethanol-sterilized razor blade (for amputation)

Temperature-controlled incubator or room (for planarian storage)

Day 1: Pre-amputation imaging and amputation

• 1
  Select a minimum of n = 10 planarians of similar size from species of choice per treatment and place them in a single 100-mm petri dish with ∼20 ml planarian water.

  Select worms that are of similar size (approximately 6 to 8 mm) and that are dark enough to allow for good contrast against the blastema when regenerating. The exact volume of water in the petri dish is not important for this step, as it is simply for storing the planarians short term.

• 2
  Fill another petri dish with 20 ml planarian water for imaging.

  Use a new petri dish for imaging. This will ensure good image quality.

• 3Use a plastic transfer pipet to move a planarian into imaging dish.
• 4
  Set magnification of the light microscope such that the planarian takes up most of the field of view but can still be kept within the frame long enough to image. Write down magnification used.

  It is recommended to keep the magnification consistent throughout the imaging session to facilitate the image analysis in Basic Protocol 2. If similarly sized planarians are used, one magnification setting should be sufficient for all specimens.

• 5
  Optimize image settings by using a black background with white light from above (Fig.1). Image each planarian pre-amputation. Save each image with clear identifiers (experimental group, worm ID number, day number, magnification; see example data in Supporting Information).

  Image the planarian near the center of the petri dish to avoid aberrations from the imaging dish. Allow the planarians to reach their gliding length before taking a picture. Resting planarians are wider and shorter, which would lead to incorrect width measurements. The planarian should be gliding in a straight line to allow for more accurate measurements. Curving of the planarian can lead to inaccurate width measurements. Avoid glare near the worm and any scratches or debris in the dish or the walls of the dish.

  To ensure that at least one high-quality image is obtained that can be used for analysis in Basic Protocol 2, it is recommended to image each planarian 2 to 3 times. Alternatively, one could quality-check the images as they are taken and repeat imaging until an image of sufficient quality is obtained.

  Keep the identity of the planarians, e.g., ID/well number, consistent across the entire experiment. This ID number should be labeled in the saved images to allow for quantification of the growth rate for each individual specimen.

• 6
  Image a scale bar or ruler using same imaging settings as for the intact planarians.

  This is required for conversion of pixels to length (mm).

  If different imaging settings are used for different planarians or different imaging days, a new scale bar image is necessary for each setting. Save the scale bar image with an appropriate name; this is especially important if using different magnification settings within one imaging session.

• 7
  Prepare two separate petri dishes (“disposal dish” and “amputation dish”) with 20 ml planarian water and one 12-well tissue culture–treated plate with 2 ml planarian water per well.

  One petri dish will serve as the “disposal dish” for the amputated pieces that will not be imaged further. Label this petri dish with the species name and the date of when amputation occurred. These pieces will be allowed to regenerate in a temperature-controlled incubator and can be re-introduced into their normal population within a week. The other petri dish serves as a temporary vessel for amputation (“amputation dish”).

  At least once during regeneration, using a transfer pipet, transfer the planarians in the disposal dish to a new petri dish filled with 20 ml planarian water to ensure a clean environment.

• 8
  Transfer a single planarian to “amputation dish” using a transfer pipet. Use an ethanol-sterilized razor blade to amputate planarian with a single transverse cut. For head regeneration studies, cut between auricles and pharynx (Fig. 2).

  If the cut is accidentally made above the auricles, make another cut. If the pharynx is cut or the cut is not straight, replace the planarian by moving it into the “disposal dish” and selecting a new worm. Image the new planarian before decapitation.

  Amputate one planarian at a time in the amputation dish to ensure the identity of each specimen is tracked throughout the experiment. Clean the razor blade following each amputation by wiping with 70% ethanol on a paper towel.

• 9
  Use a transfer pipet to remove the anterior piece and discard it in the “disposal dish”. Pipet the posterior piece into a well of the 12-well plate containing 2 ml planarian water. Transfer as little water as possible. In the case of a chemical exposure experiment, pipet the decapitated planarian into a well of the 12-well plate containing 1800 μl planarian water, making sure to transfer as little extra water as possible. Add 200 μl of 10× stock of desired exposure concentration of chemical to the well. Gently pipet up and down to mix while avoiding air bubbles.

  As an example, if using 15 mg/L (w/v) TritonX-100 (Fisher Scientific, BP151-100), CAS no. 9002-93-1) as a positive control, add 200 μl of 150 mg/L (10× stock, diluted in planarian water) to each well.

  Make sure to add chemicals within 3 hr of decapitation. It is best to add the chemicals to all the wells after imaging is complete.

  Label the 12-well plates appropriately to ensure the identity of each planarian is known throughout the experiment.

  CAUTION: Follow all state and federal guidelines regarding handling and disposal of any chemicals or chemically contaminated waste that is used. Work in a fume hood as necessary and wear appropriate PPE.

• 10Repeat steps 3 to 9 for each planarian.
• 11
  Once complete, store plates at room temperature (18° to 22°C) in the dark in a temperature-controlled incubator or room until imaging again (see steps 12 to 17).

  Note down the temperature as this can affect regeneration kinematics.

Figure 1.

Optimal imaging conditions. Examples of (A) good images for i) pre-amputation and ii) post-amputation and (B) bad images of planarians showing i) low contrast, ii) noisy background and too-low magnification, iii) a blurry/out-of-focus field of view, iv) a contracted planarian, v) a curved planarian, and vi) a blastema not within the imaging frame. Scale bars: 1 mm. Note that although color images are shown, images taken in grayscale are also suitable for this analysis.

Figure 2.

Indication of the desired amputation plane for head regeneration studies. Scale bar: 0.5 mm.

Days 4 to 7: Imaging blastema regeneration

• 12
  On day 4, prepare a clean imaging dish for each experimental group and for each control group. Fill each dish with 20 ml planarian water.

  Label the dishes appropriately so that each dish is dedicated to one exposure group. This dish will be reused for all the imaging sessions (with fresh planarian water each day).

• 13
  Use a transfer pipet to transfer a single planarian from its well in the 12-well storage plate to the imaging dish. If testing the effects of chemical exposure on regeneration, first wash all planarians by transferring them individually to new wells (keeping the same order) of a 12-well plate with fresh planarian water (2 ml/well). Transfer a minimal amount of water/chemical with planarians. For imaging, using a fresh transfer pipet, transfer each planarian individually from this “water wash” plate to the imaging dish.

  The “water wash” 12-well plate will only be used for storage of the experimental planarians during imaging. Separate transfer pipets should be used for transfer of the planarians to the “water wash” plate and for transfer into the imaging dish. Do not mix transfer pipets across chemical treatments.

  Set a timer when the planarians are removed from the chemical solution and record how long exposure was interrupted for imaging. Work quickly to minimize their time in planarian water.

  CAUTION: If working with chemicals, wear PPE and use a fume hood as appropriate for transferring worms with chemical exposure. Discard chemically contaminated solid waste as appropriate.

• 14
  Image as described in steps 4 to 6. Save each image with clear identifiers (experimental group, worm ID number, day number, magnification; see example data in Supporting Information).

  It is recommended to keep the magnification consistent throughout an imaging session (i.e., on each day) to facilitate image analysis (see Basic Protocol 2). Because the planarians are smaller post-amputation and then the blastema grows over the course of the experiment, the same magnification may not be appropriate on different days. Use a magnification that allows for the anterior region of the planarian to take up most of the field of view when at its gliding length. For imaging of the blastema, the entire planarian does not have to be within the field of view; only the blastema should (for example, see Fig. 1A). Newly regenerated planarians may not move much. Resting planarians are wider and shorter, which would lead to incorrect area measurements of the blastema. If needed, gently flush the planarian with planarian water using a transfer pipet to encourage it to elongate into its gliding length.

  Prioritize having the blastema in focus, which may cause the body of the worm to be slightly out of focus.

  As on day 1, it is recommended to image each worm 2 to 3 times or to quality-check the images as they are taken. Repeat imaging until an image of sufficient quality is obtained for each planarian.

• 15After imaging, transfer the planarian back to its original well (either in the original exposure plate or in the “water wash” plate if testing chemical exposure). Continue until all planarians have been imaged. If the planarians are kept in the “water wash” plate, continue to step 16. If the planarians are not chemically exposed and were placed back into their original 12-well plate, skip to step 17.
• 16
  For chemical exposure experiments, after all planarians in an exposure group have been imaged, transfer planarians sequentially into new wells (keeping the same order) of a 12-well plate with the appropriate concentration of chemical solution (2 ml/well) (“chemical wash plate”). Transfer a minimal amount of water with planarians. Then, transfer each planarian back into its original well in the exposure plate.

  This “chemical wash” step ensures that a minimal amount of planarian water is transferred with the planarian and that exposure conditions do not change over time when using static exposure. If using repeated-exposure conditions, replace the exposure solutions daily following each imaging session, and thus, this wash step could be removed.

  Note that this may affect the toxicity compared to static exposure without daily replacement. Save the “chemical wash” plate for consecutive days.

  CAUTION: This “chemical wash” step should be carried out wearing full PPE and in the fume hood as necessary, depending on the chemicals used.

• 17
  Repeat imaging (steps 12 to 16) on days 5 to 7. Image each planarian individually, following the guidelines provided above for the best image quality. Between imaging sessions, store the planarians in their 12-well plate in the dark in a temperature-controlled incubator or room.

  The imaging petri dish should be cleaned at the end of each imaging session (i.e., on each day). Discard the planarian water and wipe the petri dish with 70% ethanol to remove any mucus. Let it air-dry overnight and refill with fresh planarian water before imaging on the next day.

BASIC PROTOCOL 2 QUANTITATIVE ANALYSIS OF BLASTEMA SIZE WITH ImageJ

Here, the images of the planarians obtained in Basic Protocol 1 are analyzed using the free software Fiji/ImageJ (Schindelin et al., 2012). Prior to image analysis, the best image of each planarian from each imaging session should be selected and moved into an appropriately labeled subfolder for each day. Thus, there will be five subfolders (one for each imaging day), and each subfolder should contain one image per planarian. The images are individually uploaded to ImageJ and analyzed using standard techniques such as image cropping and thresholding. Each image must undergo individual contrast enhancement and thresholding. Values in pixels are converted into real length units using the image of the ruler taken in Basic Protocol 1. The planarian head width and the blastema area are measured and recorded. A template workbook is provided (see Supporting Information) for the recording and analyzing of these data. This workbook contains two sheets. The data obtained in Basic Protocol 2, as described below, will be input into the first sheet. The second sheet contains calculations and graphs where the normalized area of each blastema per day is visualized and fitted, as explained in Basic Protocol 3. It is best to analyze all images of one experiment in a single session, ensuring consistency, and to blind the analyzer to the identity of the different experimental groups, ensuring unbiased results. All relevant settings should be recorded for future reference. The protocol provides step-by-step instructions of how to conduct the analysis and is supplemented by video files that show an example analysis. Raw images for one of the control specimens are provided in the Supporting Information as an example.

Materials

Quality images of regenerating planarians (see Basic Protocol 1)

Computer with Fiji/ImageJ (version 1.53t or later, https://imagej.net/software/fiji/) installed

Google Sheets or similar software

Measuring pre-amputation head width from day 1 images

• 1For each planarian, using a computer with Fiji/ImageJ installed, choose best image(i.e., planarian is fully elongated, takes up most of the frame, is in focus, and has good contrast; see Fig. 1) from among quality images of regenerating planarians and save in appropriate subfolder for day 1.
• 2
  Open first image from day 1 and its corresponding ruler image using File > Open in ImageJ.

  Alternatively, individual images can be opened by dragging and dropping into the ImageJ toolbar.

• 3
  Convert image from pixels to length (mm) (see Video 1). Use Line tool from the ImageJ toolbar and draw a line between two mm marks (ideally choosing a larger distance for more precise measurements, e.g., between 0 mm and 10 mm) by clicking on where to start the measurement (ideally, the center of one mm mark) and then clicking on the point where to end the measurement (the center of another mm mark). Then, for each image, scale image to obtain measurements in real length units (Analyze > Set Scale). In Distance in Pixels, view number of pixels measured by the line tool. In Known Distance, enter the appropriate corresponding real length (e.g., 10 in the example) and set the Unit of Length as the appropriate unit (e.g., mm in the example). If all images have the same scale, check box for Global. To apply the scale to the selected image, click OK.

  This step will allow for measurements to be obtained in actual units (i.e., mm) rather than pixels so that results can be compared across different imaging conditions. By checking Global, this scale will be applied to all the images that are uploaded to ImageJ during this session (until ImageJ is restarted or the scale is removed). The images will now display the length as in “mm” instead of “pixels.” Only select Global if all images were taken at the same magnification. If images have different magnifications, this step will need to be repeated for each image.

• 4
  Measure the width of the planarian head (see Video 2):

  1. Zoom in around the head of the planarian to increase accuracy. Place cursor on region of interest (head) and use either up and down arrow keys or “+” and “-” keys to zoom in or out.
  2. Select Line tool from the ImageJ toolbar. Create a straight line by clicking on opposite edges of the worm, slightly below the auricles (Fig. 3A).

     The width is slightly larger right below the auricles and then becomes fairly constant for the remainder of the pre-pharyngeal region. Because the cut is performed in this central pre-pharyngeal region, this constant width should be used as reference.

  3. Obtain a measurement of the width by clicking Ctrl+M.
• 5
  Open workbook provided in the Supporting Information in Google Sheets or similar software.Navigate to Data sheet. Fill in the respective information in the appropriate sections:

  1. Enter the name of the worm under Worm ID.
  2. Copy the measurement under Length from the Results window into the workbook under Day 1 Width.
• 6
  Repeat for remaining planarians.

  All planarians should be treated the same in the analysis pipeline.

Video 1.

Download video file ^{(44.4MB, mov)}

Tutorial on how to set the scale of an image in ImageJ.

Video 2.

Download video file ^{(36.3MB, mov)}

Tutorial on how to measure the width of a planarian on day 1 in ImageJ.

Figure 3.

Overview of key image analysis steps in Fiji/ImageJ. (A) Example of day 1 head-width measurements: i) The original image, with the head region boxed. ii) Zoomed-in view of the boxed region from i. The white line indicates where the width should be measured. (B) Example of an image during regeneration, with comparison of the i) original image and ii) the image after converting to 8-bit and performing Enhance Contrast. The blastema region is boxed. Scale bars for (A) and (B): 0.5 mm. (C) Analysis of the blastema region: i) duplicated boxed region from ii, ii) the binarized image after thresholding, and iii) the outline of the analyzed area (1). (D) Alternative method to calculate the blastema area using the Polygon tool.

Blastema area analysis using images from days 4 to 7

• 7For each planarian, choose the best image (i.e., planarian is fully elongated, takes up most of the frame, is in focus, and has good contrast; see Fig. 1) for each day of regeneration. Save in a subfolder for the specific day, as for the day 1 images.
• 8Upload the best image for a specific planarian and day into ImageJ, as instructed in step 2 for the day 1 images.
• 9
  Following step 3, set the scale for the image to obtain area measurements in real units (see Video 1).

  As applicable, this scale can be applied to all images using Global or should be repeated for individual images (as outlined in step 3).

• 10
  If necessary, convert color images into 8-bit grayscale (Image > Type > 8-bit).

  Color images/cameras are not required for this analysis but can be used.

  An example of how to perform steps 10 to 15 can be found in Video 3.

• 11
  Enhance the contrast between light and dark pixels (Process > Enhance Contrast). Input 0.15% for Saturated Pixels. Leave all boxes unchecked.

  The Enhance Contrast feature increases the contrast in the image based on the allowed number of pixels indicated by the percentage of saturated pixels and helps to distinguish the blastema. The input for saturated pixels depends on the imaging conditions and needs to be determined empirically. Adjust this value if the suggested value does not work. Greater values increase the contrast more.

• 12
  Isolate the blastema from the rest of the planarian body:

  1. Draw a tight box around the blastema using the Rectangle tool in the ImageJ toolbar by clicking and dragging the mouse horizontally. Adjust the box size by dragging the small, white squares on the box itself.
  2. Right click over the box and select Duplicate. In the pop-up window, change the title of the image if desired. Ensure Ignore Selection is unchecked. Click OK.

     A new image of just the region of interest will open.

     It is important to make the box as tight around the blastema as possible, especially making sure to not capture too much of the body of the worm (Fig. 3B and 3C) while not losing any parts of the blastema. This will reduce the number of irrelevant pixels that get incorporated into the blastema analysis. If the planarian is at an angle, it may be helpful to rotate the image (Image > Transform > Rotate) or use the Rotated Rectangle tool (long click on the Rectangle icon in the ImageJ toolbar and select Rotated Rectangle) to get an adequately tight box.

• 13
  Binarize the newly opened image of the blastema region of interest by applying a threshold (Image > Adjust > Threshold):

  1. Check boxes for Don’t Reset Range and Dark Background.
  2. In the drop-down menu on the left, make sure thresholding method is set to Default to separate the planarian from the background.
  3. In the drop-down menu on the right, keep display of the image set to Red.
  4. Adjust minimum threshold with the upper slider and maximum threshold with the lower slider until the blastema is isolated from the rest of the planarian body (Fig. 3C). Ensure that the blastema is clearly outlined in red without any holes. Compare to the original image underneath the red coloring to determine whether the thresholding is adequately picking up the blastema.
  5. Press Apply when satisfied with the settings.

     The Log window may pop up and display “Black background not set in Process>Binary>Options; inverting LUT.” This can be ignored.

• 14
  To specify the measurements for the area analysis, click Analyze > Set Measurements. Check Area and click OK.

  Other measurements, such as length, can also be selected if desired. This step only needs to be performed once per ImageJ session. This setting will be saved for the subsequent images.

• 15
  Calculate the blastema area for the binarized image using Analyze > Analyze Particles:

  1. Set area range in Size (mmˆ2) to 0-Infinity.
  2. Use the drop-down menu in Show and select Outlines to visualize the area being computed. Check boxes for Display Results, Clear Results, Include Holes, and Overlay.

     A new window (Drawing) will open to show an outline of the areas being calculated (Fig. 3C, iii).

     The Results window lists the areas that have been identified and calculated within the specified region of interest. The largest area corresponds to the blastema. Visually verify that the number for the largest area corresponds to the blastema in the Drawing window.

     Particles (objects) outside of the area range provided in Size (mmˆ2) will be ignored. Start by allowing all particles to be identified (range of 0-Infinity). If the range of blastema sizes is known, a lower cut-off value can be added for future analyses to ignore extraneous particles, which can be helpful but is not necessary. For example, in our dataset, a lower limit of 0.01 mm² (Pixel Units unchecked) was sufficient to differentiate the blastema area from other particles.

• 16
  If adequate thresholding cannot be achieved in steps 13 to 15 to isolate the blastemal from the planarian body due to low contrast in the images, isolate the blastema area manually using the Polygon tool in the ImageJ toolbar:

  1. Draw a polygon around the blastema by clicking where each vertex should be placed until a closed shape is formed (Fig. 3D).
  2. Right click over the outlined blastema and select Measure.

     The area of this region will be listed in the Results window.

• 17Copy the value corresponding to the blastema area from the Results window into the workbook provided in the Supporting Information. Match data to the correct Worm ID and copy the blastema area for the respective day under Raw Blastema Area.
• 18
  Close all windows and repeat analysis for the remaining planarians and days.

  It is recommended to save the binarized image and Drawing window for quality-control purposes to verify that the area was measured correctly should any issues with downstream analysis occur.

Video 3.

Download video file ^{(157MB, mov)}

Tutorial on how to measure the blastema area in ImageJ.

BASIC PROTOCOL 3 QUANTIFICATION OF BLASTEMA GROWTH RATE

The data obtained in Basic Protocol 2 for the blastema area as a function of time are compiled and analyzed in a spreadsheet to calculate the blastema growth rate. The blastema area is first normalized by the squared width of the original planarian. The normalized blastema areas over time for individual planarians are fitted by a linear best-fit curve using a template workbook provided in the Supporting Information. The protocol below shows how the blastema growth rate would be calculated from the width and blastema area measurements obtained in Basic Protocol 2 and how these measurements can be statistically compared across experimental groups.

Materials

Google Sheets or similar software

Data for Day 1 Width and Raw Blastema Area (see Basic Protocol 2)

1. Open workbook provided in the Supporting Information in Google Sheets or similar software to view all data for Day 1 Width and Raw Blastema Area for each planarian in the Data sheet.

2. Calculate Normalized Blastema Area for each day by dividing the area for each day by the Day 1 Width squared.

   For example, the normalized area for day 4 would be =C3/$B$3ˆ2, where the duplicated number indicates the row number (here, row 3). The $ indicates to use the exact cell position.

   The provided workbook is pre-filled with this calculation so that the data will automatically update when the raw width and area values are inputted. When adding more data, select cells in columns G to J and drag the blue box in the bottom right corner to add the commands into subsequent rows in these columns.

3. Navigate to the Graphs sheet.

   The Normalized Blastema Area for each day from Data should import to this sheet automatically.

   As necessary, drag down the cells to extend the commands down the columns, as explained in step 2.

4. To visualize the results, create a scatter plot of Normalized Area vs Days for each experimental group, with a separate line for each planarian:

   1. Highlight cells B2:E3.
   2. Go to Insert and then Chart.
   3. Double click on the chart produced and edit Setup menu as follows:

      1. Under Chart type, select Scatter chart.
      2. Under Data range, add B2:EX, where X is row number of the last worm in the dataset. iii. Check Switch row/columns and Use row 2 as labels.

         This will allow the data to be input as rows rather than columns and will automatically provide the correct data labels for the x-axis and Series.

   4. To add data from more planarians from the same experimental group, click on Add Series and add cells corresponding to the Normalized Blastema Area for the respective planarian, with each planarian as its own series.
   5. Change the name of the graph as desired (such as to the experimental group name) under Customize > Chart & axis titles > Title text.

      A scatter plot will be generated with each planarian as its own series (different color).

5. Visualize linear best-fit line for each planarian:

   1. Click on a data point for the planarian of interest to select its data.
   2. In the Chart Editor, navigate to the Customize tab.
   3. Under Series, check Trendline. Select Type as Linear.
   4. Repeat for each planarian by selecting a data point from that planarian’s series and then repeating steps 5a to 5c.

      The scatter plot with the best-fit line is used to visualize the data. This can be used to qualitatively compare across the groups and to check for any outliers between individual data points or across all data points for a particular planarian.

6. Repeat steps 4 and 5 for each experimental group.

7. Under Slope, calculate the slope of the best-fit line for each planarian using =LINEST( [known_data_y], [known_data_x]), where known_data_y should incorporate the normalized blastema area from days 4 to 7 for the planarian (i.e., BX:EX, where x is the row number) and known_data_x are the day numbers, corresponding to cells B2:E2.

   LINEST will calculate a linear trend of the data using the least-squares method. The slope will be output in column F, and the y-intercept will be output in column G. Only the slope will be used for downstream analyses.

8. Calculate the average growth rate and associated standard deviation for each experimental group:

   1. Under Average Growth Rate, use =AVERAGE (range). Change range to include the slopes for all planarians in an experimental group.
   2. Under SD, calculate the standard deviation as =STDEV (range). Use the same range as in step 8a.
   3. Repeat for each experimental group.

9. Perform a Student’s t-test to determine whether a statistically significant difference exists between the groups. In column K (T-test), calculate p-value using =TTEST(range1, range2, tails, type), where range1 should incorporate the slopes for the control group and range2 is the cells containing the slopes for the experimental group. Set tails to 2 to evaluate both increases and decreases in regeneration rate. Set type to 2 [a two-sample equal variance (homoscedastic) test] or 3 [a two-sample unequal variance (heteroscedastic) test], as appropriate.

COMMENTARY

Background Information

Planarian regeneration has long been a fascinating and accessible phenomenon to study, dating back to early studies by George Shaw in 1790 and Thomas Hunt Morgan in 1889 (Morgan, 1889). Studies of regeneration have typically relied on qualitative observations of gross features such as head shape and regeneration of the eyes and auricles. Although these types of measures are still in use today, some researchers have also begun to develop quantitative measures of regeneration to allow for the detection of subtler defects that may arise from developmentally toxic chemicals. Manual measurements of blastema length at different regeneration time points have been used to measure regeneration kinematics (Córdova López et al., 2019; Rodrigues Macêdo et al., 2019). Notably, this technique does not incorporate any measures to scale by the original size of the planarian. Similarly, measures of blastema area (with and without normalization by the original size of the planarian) have been used (Balestrini et al., 2014; Kustov, Tiras, Al-Abed, Golovina, & Ananyan, 2014). Prior studies only examined regeneration on one day (day 3 or day 7) and thus did not incorporate regeneration kinematics. In contrast to these methods, the set of protocols described here quantifies the rate of regeneration across days 4 to 7, which allows for the detection of both gross defects that could be captured on a single day (as in the previous methods) and defects in growth rate. Importantly, here, the area of the blastema is normalized by the squared width of the planarian. We have previously found that this is the most robust way to normalize by planarian size, and we have shown that this method can detect regeneration defects due to chemical exposure (Hagstrom, Cochet-Escartin, Zhang, Khuu, & Collins, 2015).

Although the protocols described here are focused on studying the effects of chemicals on head regeneration, the technique is broadly applicable to any situation where one would want to compare regenerative capacity, such as after gene knockdown using RNAi (as in Hagstrom et al., 2018) or under different environmental conditions (e.g., temperature), as in the classic work by C. M. Child (Child, 1911). Moreover, this method could be applied to study any regenerative blastema, such as in tail-regenerating head fragments. This set of protocols is appropriate to study conditions where one desires to investigate effects on stem cell dynamics (proliferation, migration, differentiation) and patterning, all processes relevant to development. A variety of toxicant classes, including solvents, disinfectants, pesticides, and silver nanoparticles, have been found to affect blastema regeneration (Córdova López et al., 2019; Hagstrom et al., 2015; Kustov et al., 2014; Rodrigues Macêdo et al., 2019). Moreover, a lack of defects in blastema regeneration can be an equally important observation. For example, exposure to the pyrethroid insecticide permethrin (Hagstrom et al., 2015) and the alkaloid berberine (Balestrini et al., 2014) were found to impair eye regeneration in the absence of significant effects on blastema growth. These findings suggest that these compounds may have specific neurotoxic effects on the visual system rather than general developmental toxicity.

Critical Parameters

The first step in ensuring robust measures of blastema size (Basic Protocol 2) is to obtain high-quality images (Basic Protocol 1). The images should be of high contrast, allowing for the distinction of the unpigmented blastema from the dark planarian body and the background. Use new petri dishes to avoid noise due to dust or scratches. When possible, the planarians should be at their gliding length and not overly stretched or contracted. This is important, as we only have access to measures of two dimensions (width and length). To streamline the image analysis, it is critical to standardize the imaging conditions across all samples, including lighting and contrast. Note that an alternative approach to manually isolate the blastema is provided, which can be used if sufficient contrast cannot be achieved to get robust binarized images. This may be needed during later stages of regeneration when the blastema regains some pigment. Although the analysis does account for different worm sizes, it is prudent to use planarians of similar size to eliminate size as a confounding factor in the regeneration kinematics. When handling the planarians, be careful to not damage them. Any planarians that get damaged during the experiment should be discarded from the analysis.

Handle all exposure chemicals according to manufacturer and regulatory guidelines. Use a fume hood as necessary. Imaging can be done in untreated planarian water (e.g., 1× IO or 1× Montjuic Salts), but planarians should be transferred with care back to their chemical exposure solutions after imaging. Chemically contaminated pipets and petri dishes must be disposed of according to regulatory guidelines.

As some of the steps described could be subject to researcher-to-researcher variability, it can be helpful to analyze multiple images (i.e., similar to technical replicates) using two independent analyzers each, as was done previously (Hagstrom et al., 2015). This allows for the data to be averaged, providing more robust measures of blastema size with their associated error terms.

Troubleshooting

Table 1 describes some common problems that may be encountered when executing these protocols and offers possible causes and solutions.

Table 1.

Troubleshooting Guide for Quantitative Blastema Growth Analysis

Problem	Possible cause	Solution
Difficulty to keep planarian in the field of view (FOV)	Planarian moves in reaction to the light	Lower the light intensity and/or lower the magnification to create a larger FOV. Try to keep the planarian as large as possible in the FOV to maximize resolution.
Regenerating planarian is contracted	As tail fragments regenerate, they are less mobile and thus more likely to be contracted	Pipet some planarian water onto the tail fragment or agitate it gently by moving or tapping the dish to get it to elongate.
Background inhomogeneous	Background is not perfectly smooth, and any texture is visible at higher magnification	Use a lid of a clean petri dish underneath the dish to be imaged to separate the focal planes. This will blur the background in the image and thus make it appear more homogeneous.
Image of low quality (blurry or low contrast)	Image conditions have not been optimized	Improve image conditions by changing the lighting and adjusting the focus and any mirrors (as applicable). Re-image the worms if possible.
Poor thresholding	Original thresholding produces holes within blastema area	Change the thresholding method before using the sliders to adjust the maximum and minimum threshold values. Try to fill in the holes using Process > Binary > Fill Holes. Use the Polygon tool to manually define the area of the blastema.
Blastema area is not correctly identified in Drawing output	Background could be identified instead of the blastema	Try unchecking “Dark Background” during the Threshold step to invert the binary image. Then, try Analyze particles again. If necessary, revert to the original image and prepare for image analysis again. Alternatively, use the Polygon tool to manually define the area of the blastema.
Data analysis not working with spreadsheet provided	Incorrect cells are being referenced or wrong data were added to the cells	Start with the original copy of the spreadsheet and repeat the data entry and analysis.

Statistical Analysis

The normalized blastema growth rate (1/day) is calculated as the slope of the linear regression of the normalized blastema area over time for each planarian. Blastema growth rates are expected to be normally distributed. Thus, a two-tailed t-test is an appropriate method for comparing blastema growth rates between exposure groups. If multiple groups are processed at once, ANOVA can be conducted. A template for the analysis is provided in the Supporting Information.

Understanding Results

This set of protocols will yield blastema growth rates for regenerating planarians using basic light microscopy images. Images shown in the figures were acquired on an inexpensive digital microscope (Andonstar 5-inch Screen 1080P Digital Microscope, Amazon) in a teaching lab environment. To obtain robust quantification of blastema growth, the images of regenerating planarians acquired in Basic Protocol 1 must be of good quality. Figure 1 shows what images are suited or not suited to further analysis. The head region is of special importance, as this is what is used in the analysis in Basic Protocol 2. If other parts of the planarian are slightly out of focus or curved, that is acceptable. The background should be as homogenous and have as much contrast relative to the planarian and blastema as possible.

Figure 2 shows where amputation needs to take place when studying head regeneration, as we have done in Basic Protocol 1. Figure 3 provides an overview of the key steps of the image analysis conducted in ImageJ to determine the blastema area during regeneration (see Basic Protocol 2). The images are thresholded to separate the blastema from the background and the planarian body to allow for a measurement of its area. The blastema area is computed using a built-in particle analysis algorithm and visualized to allow for manual quality control of the measurement. Although we use D. japonica planarians, the protocols can be used without modification for other planarian species, such as the commonly used S. mediterranea or G. dorotocephala. For all three species, the unpigmented blastema tissue can easily be distinguished from the remaining pigmented body. Thus, for any dark pigmented planarian, the protocols can be applied without modifications. We illustrate and explain possible results that may be obtained for a chemical exposure experiment using these protocols for the species D. japonica. Ten D. japonica planarians were each exposed to 15 mg/L (w/v) TritonX-100 (Fisher Scientific, BP151–100, CAS no. 9002–93-1) in IO water or to IO water only (control) within 3 hr of amputation. This concentration of TritonX-100 has previously been found to induce regeneration delays (Hagstrom et al., 2015). A sample image sequence of one representative member of each exposure group (Fig. 4A) and quantification of the blastema growth rate (Fig. 4B to 4D) are shown. In agreement with our previous results, planarians exposed to 15 mg/L TritonX-100 had a significantly decreased blastema growth rate compared to the control group (p = 0.07, two-tailed Student’s t-test), indicating a reduction of blastema regeneration in the chemically exposed planarians. The raw images and analysis for these data are provided as example data in the Supporting Information.

Figure 4.

TritonX-100 (15 mg/L) induces regeneration delays in D. japonica planarians. (A) Representative images of the regeneration time course of control planarians (top) and planarians exposed to 15 mg/L TritonX-100 (bottom). Scale bars: 0.5 mm. (B and C) Growth curves for control planarians (B) and planarians exposed to 15 mg/L TritonX-100 (C). Points indicate the normalized blastema area, and lines indicate the linear regression for each individual planarian (different colors). (D) Comparison of the average control and chemically exposed growth rates using a two-tailed Student’s t-test. * indicates p < 0.05. Error bars indicate the standard deviation for n = 10 planarians per group.

Time Considerations

Imaging blastema size (Basic Protocol 1) will take 1 week. The imaging process will take about 30 to 60 min per 10 worms, depending on experience level. Image analysis (Basic Protocol 2) takes ∼5 min per worm. Quantification of the blastema growth (Basic Protocol 3) rates takes ∼5 min/group.

Data Availability Statement

The data that support the findings of this article are available in the Supporting Information.