The size and shape of activated sludge particles is critical for wastewater treatment efficiency. Yet their measurement is often ad hoc. This protocol provides a repeatable, well defined, and semi-automatable measure for their measurement.
The main advantage of this protocol is that it can measure a large population of particles over a wide range of morphologies while reducing subjective and systematic bias. While this technique is focused on activated sludge, technically anything that has the same particle morphology such as microplastics can be measured with this method. When attempting this technique for the first time, try making one plate at a time and practice the physical motions of making that one plate.
Also, take your time with the microscopy. The video version of this article is important because sampling and plate preparation require a lot of little techniques that are best communicated visually. Also, showing and seeing good pictures of plates are important to ensure reproducibility.
Demonstrating this protocol today is Joseph Weaver, a PHD candidate in my lab. To begin, acquire a representative sample from a well mixed portion of the reactor. Gently mix the sample and then immediately pour the determined sample volume into a 15 milliliter centrifuge tube.
Next add five microliters of one percent methylene blue to each sample. Cap and mix the sample by gently inverting the tube at least three times. Allow the samples to stain for at least five but no more than 30 minutes at room temperature.
Once stained, transfer a sufficient volume of melted 7.5 percent agar and deionized water to the centrifuge tube in order to bring the total tube volume to between 6.5 and 9 milliliters. Then recap the centrifuge tubes and gently invert them at least three times to mix the sample into the agar. While pointing the cap away from oneself, open the cap.
Pour the tube contents of each tube into their own 100 milliliter plastic petri dish while gently rocking the dish to achieve a full, smooth coating and a visually uniform distribution of particles. Allow the plates to cool at room temperature for at least 5 minutes, until the agar solidifies. Place the uncovered plate face up on the microscope stage of a stereo microscope that is capable of 10 to 20 times magnification.
Illuminate the sample from below with even diffuse light, using equipment such as an LED illuminator stand or light plate. Open the image capture software and ensure the microscope light path is set to photo. Then, select the appropriate camera from the camera list.
Adjust the microscope so that multiple particles appear in the focal plane with large, well defined edges. Use a magnification of 10 to 20 times to measure particles while maintaining a relatively deep focal plane. Temporarily remove the agar plate and place the micrometer on the stage.
Adjust the fine focus until the graduations on the micrometer appear sharply focused in the image capture software. And calibrate the pixel to micron ratio. Next, set the zoom to 100 percent by clicking on view and selecting actual size.
Then select options and go to calibrate. Here, align the red calibration bar in the main viewport along the long axis of the micrometer. With the vertical bars centered on the zero and 200 micron graduations.
In the calibrate dialogue box, enter the current magnification level and actual length of 200 microns. If already calibrated, select magnification from the menu bar and then select the current magnification level to confirm the calibration. In the measurements menu, go to line and select arbitrary line.
Click on the intersection of the zero graduation and long axis of the micrometer. Then click again on the intersection at 200 micron line and the long axis. A correct calibration should display as approximately 200 microns.
Delete the line by clicking the select object button, clicking on the line, pressing delete, and pressing yes on the confirmation box. Next, replace the agar plate and adjust the fine focus to achieve maximum detail in the imaging software. To accomplish this, first increase the bit depth of the maximum value allowed by selecting the radio button in the bit depth panel of the camera sidebar.
Also, set the software to acquire grayscale images by selecting the appropriate radio button in the color grade panel of the camera sidebar. Now, collapse any open sidebar panels between exposure and histogram. Reduce the gain to one and increase the exposure until a clear image appears in the viewport and until the histogram appears as a distribution that is not clipped by either end of the histogram box.
Adjust the histogram to avoid over and under exposure. In the histogram panel of the camera sidebar, slide the left boundary of the histogram to just outside the lowest values and the right boundary to just outside the highest values. Save the image as an uncompressed TIF file.
Include magnification information in the image metadata by checking the save with calibration information box when saving. Select another area that does not overlap the previous images by following a path which alternates between left to right and right to left as one moves down the plate. Capture at least 500 visually estimated particles for analysis.
Acquire the particle analysis code by cloning the GIT repository. At the command line, retrieve the latest version of the code by typing in the command shown here. Install the analysis code following the instructions in the read me text file found in the top level directory of the cloned repository.
Next, edit a text file listing the directories to be processed along with optional parameters. Refer to the examples and analysis subdirectory for a list of parameters and examples. Now, run the analysis on the command line by typing the command shown here.
Fiji path is the directory in which imageJ-win64. exe is located. And paramsfile the location of the text file describing the analysis setup.
The name of the executable may differ depending on which operating system Fiji is installed on. The analysis will run automatically and take a few minutes to a few hours depending on the size and number of images. Next, examine the quality control files located in the overlay subdirectory of the specified output directory.
Note images with spurious mist or poorly captured particles, all apparent as shaded outlines which did not match the background. That's it. The data are now ready for experiment specific analysis and figure generation.
Although no particle thresholding method is perfect, here is an example of acceptable results. When evaluating QC images, there are three common errors found. Failure to accurately conform to the particle boundaries, failure to identify particles, and artifact inclusion.
Here, the particle distributions between two experimental reactors over time are displayed and combined with qualitative metadata noted by the researcher. This chart shows that in this case, the reactors had generally similar particle size distributions which tended towards a larger mean and wider spread as time progressed. When performing this procedure, it is critical to evenly coat the plate with a thin layer of agar containing uniformly distributed particles.
Be sure to practice and take your time. Beyond this procedure, interesting particles could be excised from the agar and studied. In terms of data, the resulting files from this protocol are well defined and the sky's the limit analytically.
This technique paved the way into providing new insights into aerobic granular sludge. It's morphology is very important and it allowed us to measure it in a standard way. Wastewater is biologically active.
Bio safety level one practices are encouraged. Also, how agar can bubble over when microwaved and can spray hot droplets when opening the centrifuge tube.
