Leaf spot patch analysis caused by Blackleg

My research involves the classification of a leaf spots on turnip derived from the pathogen blackleg. I had hypothesized that spatial patterns of pixels based on image classification are related to manually classified spatial patterns of observed disease on turnip leaves because disease has a characteristic spectral signature of infection on the leaves. This post focuses on the analysis of the clusters based on image classification through segmentation as well as the manually classified clusters. These clusters of pixels are expected to be representative of the diseased patches on the leaves. Here we seek to obtain some patch statistics which will be compared for relationships and accuracy at a later time. A large portion of this process went into image processing before the analysis could be conducted. All of the image processing took place in ArcGIS Pro. The next step involved the patch analysis which was conducted in Fragstats.

Some questions I asked myself about element A & B of my hypothesis included:

Can diseased patches be successfully identified and isolated by computer image classification and through manual classification?

If yes; what is the area and perimeter of the diseased patches based on image classification and manually classified diseased patches? I was mostly looking to obtain and gain some experience in the patch analysis provided by Fragstats. Much more thorough analysis can be and will be completed in Fragstats when variable A & B are compared for accuracy assessment in exercise two.

Tools used and methodology
The image processing and classification of pixels was conducted in ArcGIS Pro. This analysis required extracting both manually classified diseased patches and computer classified patches. I began by uploading band 3, which captures light energy in the red wavelength at 670 – 675 nm.

1. For computer classification of the image I used Segmentation.

a) I went to the Imagery tab and the Image Classification group. Click the drop-down arrow and an option for Segmentation should appear. Be sure the layer you desire to perform the segmentation on was selected prior to selecting the Segmentation Classification Tools. In the pane you have options to adjust Spectral detail, Spatial detail and Minimum segment size in pixels. There are variable and depend on image resolution, level of accuracy you wish to achieve, etc. Because my resolution to high I chose for higher spectral detail and spatial detail. I adjusted the spectral detail from 15.50 to 17 and the spatial detail from 15 to 17 as well. For the minimum segment size in pixels I chose 10. As mentioned these values are variable and although the standard values worked well for segmentation, I previewed other values and achieved greater results by adjusting. After previewing other options and deciding what values work best, click the Run button in the bottom right corner of the pane.

b) Masking all the cells that weren’t diseased was the next step and crucial to this diseased region selection process. To do this, go to the Analysis tab and the Raster group to find the Raster Functions. Open the Raster Functions and it should appear in the pane to the right. Search Mask and select it. Open the segmented image in the raster box and three options will be available for Included Ranges. These three options are representative of the blue, green and red bands. Because the data was adjusted from 12-bit to 8-bit in the segmentation process, we should have a range of values between 1 and 255. Despite having three band options, all the bands hold the same value because we are only working with the red band. Click on the segments you wish to include in the working window to check their values. Include a minimum and maximum which should be the same for all 1, 2 and 3 that includes the segments you want. The range I had included was 100 to 160, which is the RGB.Pixel Values that appear when clicking a pixel. Once you have these entered, click create new layer in the bottom of the right pane.

c) Next, I used the Clip function by navigating to the Imagery tab, the Analysis group and selecting Raster Functions. In the window pane to the left a search bar can be found along the top where you can enter Clip. Although there are other methods and locations the clip function can be found, I experienced different results depending on how I navigated to clipping, as well different clipping options in the window pane. In the Parameters section for Clip Properties, select the drop-down arrow for the raster you previously segmented. Leave Clipping Type and Clipping Geometry/Raster as the default. Adjust the active map view by zooming in or out to the region you want preserved after the clip. I use as small of area as I possibly can when clipping, while keeping all required segmented regions. Click the Capture Current Map Extent button found just to the right of the extent options in the pane (Green square map). To clip, click Create new layer at the bottom of the pane and a new map layer with the clipped region should appear in the map contents.

d) From here, go to the Analysis tab and Geoprocessing group where you can find the Tools. Click the Tools to open the options in the left pane. Here we want to use the Raster to Polygon (Conversion Tools) which can be found by searching at the top of the tools window pane. Use the most recent segmented-clipped raster for the Input raster, leave the Field blank and select where you would like the Output Polygon features to save. I left the remaining option as the default and clicked the Run arrow in the bottom right corner. Although we need the final product to be in raster format, this step is important for creating separate polygons which are grouped together as one unit.

e) As mentioned we must get the polygons back into a raster. Before we use the Polygon to Raster tool, polygons which overlap must be merged. This is one of the two reasons why we converted the raster to polygons. When the segmentation was conducted in (step d), diseased regions were not all categorized in the same bin. This resulted in regions which were maintained after masking but are different shades grey. Even one diseased patch may have two to three tones to it, resulting in segments for a patch. Now that everything is a polygon, we can merge these polygons that should be one polygon. Click the Edit tab and select Modify in the Features group. In the Modify Features, scroll down to the Construct group and click Merge. In Existing Features click Change the Selection and while holding shift, select the polygons which belong in one group. They will be highlighted in blue and appear in the pane if properly selected and if so, click Merge. Navigate to the Map tab and Selection group and click Clear. Use the same merge steps to merge any other polygons which belong to the same diseased lesion but were classified separately in raster segmentation.

f) Finally, the polygons can be converted back to a raster for the last step of image processing. Click the Analysis tab and find the Geoprocessing group to Tools once again. Search Polygon to Raster (Conversion Tools) in the Tools pane and select it. Use the polygons layer for Input Features and the window pane options should adjust to accommodate your raster. The only adjustment I made was to the Cellsize which I changed to 1 from 0.16 to maintain the cell size in my original raster. Select Run in the bottom right corner of the pane for the final layer. Each of the polygons should now be converted to a rasterized polygon with a unique ObjectID, Value and Count which can be found by going to the Attribute table for that layer or clicking on a group of pixels. This allows for each diseased patch to be aggregated and analyzed as such in Fragstats.

g) Go to the final raster layer, right click and go to data and export data. What I wanted is a TIFF file for analysis in Fragstats.

2. To compare the accuracy of the segmentation as a classification method for diseased pixels, manual classification was used as a ground truth technique.

a) For manual classification of the original red band, the Training Sample Manager was used. This allows for manual classification which can be used for supervised machine learning techniques of classification. Here, I simply used it to select diseased regions manually, but have an end goal of using the support vector machine learning model.

b) To get to the Training Sample Manager, go to the Imagery tab and Image Classification group and select the Classification Tools where you can click Training Sample Manager. In the Image Classification window pane go to create new scheme. Right click on New Scheme and Add New Class and name it diseased pixels and supply a value which is arbitrary and a description if desired. Click ok and select the diseased pixels scheme and then the Freehand tool in the pane. Draw an outline around the diseased regions in the image. Save these polygons and the classification scheme. Go to Add data in the Layer group on the Map tab to upload the training sample polygons.

c) Once the polygons are uploaded, use the Polygon to Raster protocol which can be found in step 1, part (f) followed by part (g).

Fragstats was used for the analysis of the polygons. For exercise 1 the intent in Fragstats was to simply obtain information about the diseased patches that were extracted from the red band image. This included the pixel number, area, perimeter, perimeter-area ratio and shape index. Many more options were available for patch analyses and will be considered for exercise 2.

1. Open up Fragstats for the patch analysis.

a) To import your first image click Add layer and go down to GeoTIFF grid and select it. In the Dataset name, search for your tiff file you created in ArcGIS Pro and select it and click ok. You have to do this for each of the datasets. Then go to the Analysis parameters and click Patch metrics in the Sampling strategy. For General options hit Browse to find a location for the output to save and click the Automatically save results checkbox.

b) Click on the red box called Patch metrics and in the Area – Edge tab select the Patch Area and Patch Perimeter. Click on the Shape tab and click Perimeter – Area ratio and Shape Index.

c) In the upper left corner, you can hit Run and it will check everything you have uploaded and the analysis you have selected. You must then hit Proceed after it checked the model consistency and it runs the analyses.

d) After this hit the Results options for viewing the output.

The patches aligned pretty well through visualization in ArcGIS Pro when comparing the two layers. The comparison between patches will be done in exercise 2. I did notice that the segmentation process missed many of the smaller diseased patches. Looking at Image 2, you can see that they were segmented from the surrounding regions but were lumped into a bin with that caught areas that appeared lighter in the image because of a reflection. Different thresholds could be used in the segmentation process in order to include those smaller patches. This could maybe be done if they followed a parameter involving shape. This may be considered for future segmentation. A concern is that you set the threshold too low and it includes many regions that aren’t diseased but include all the diseased regions also. This would result in many false positives for disease. The alternative is to set a high threshold value for diseased regions which would result in false negatives. The idea is to find a good balance between the two. Currently the segmentation is strictly based on the segments value which is attributed to the reflectance in the red band. As mentioned another parameter worth considering is shape. Since the diseased regions tend to be leaf spots resulting in a circular area typically, adjustments could be made to include more circular patches that don’t extend past a certain number of pixels. In table 1 we see that number of pixels for patches ranges from 8 to 50. The patch analysis provides some insights into possible spatial factors that explain these segmented regions and how the classification process could be done more accurately.

Table 1. Showing the 11 different patches that were manually selected and analyzed in Fragstats. The color indicates the corresponding patch that was detected through segmentation shown in Table 2.

Table 2. The 4 different patches that were identified through segmentation and analyzed in Fragstats. The color indicates the corresponding patch that was detected through segmentation shown in Table 1.


Critique of the method
The MicaSense red-edge camera has 5 bands which can be very helpful for applying different vegetative indices and compositing bands in order to help bring out the variation in spectral signatures between diseased and un-diseased tissue. Although the pixel size is just under 1 mm, which appears to be adequate for identifying lesions on the leaf, the bands do not have perfect overlap. This is due to the design of the camera which has five different lenses for the five different bands that are all separated by one to two inches. This could be corrected for if the extent for each of the five bands was manually adjusted for a near perfect overlap. Until this is resolved, the red band seems to show the most variation in pixel value for diseased patches in comparison to the other four bands and was used for this analysis.

Additionally, the amount of processing that is done in part 1 step (a) to get the segmented raster patches is has many steps. Methods for speeding up this process need to be considered for future analysis especially when conducting this analysis on the 500 leaves.

As mentioned before, Fragstats has many more statistically capabilities that were not applied in this analysis. Getting more statistics on the manually and segmented patches would be helpful for determining the level of accuracy as well as other parameters worth considering.


Image 1. The red band tif file uploaded into ArcGIS Pro for classification. The white patches are what we would expect the diseased regions to look like.

Image 2. The result of the segmentation performed on Image 1 described in step 1 part (a)

Image 3. The patches determined based on segmentation in part 1 from Image 2. Separate patches were created by using the Raster to Polygon tool, followed by the Polygon to Raster tool. Two layers were turned on here for the intent of showing the overlap between the raster patches and the original image.

Image 4. The patches determined based on the manual classification described in part 2 from Image 1. Training sample manager followed by polygon to raster was used to get these patches. Two layers were turned on here for the intent of showing the overlap between the raster patches and the original image.

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1 thought on “Leaf spot patch analysis caused by Blackleg

  1. jonesju

    Taylor, very complete account of your work. It appears that the time-consuming classification is able to detect only a subset of the large, manually identified patches. For Ex 2, it would be useful to conduct an error analysis and determine whether the patches that were missed by the classifier have other characteristics (besides being small) that might explain why they weren’t captured by the classifier. Maybe in Ex 3 you might try a much simpler automated classifier, and also use a training dataset, and consider introducing spatial constraints into the classifier.

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