Team:Lambert GA/Hardware

MICROMESH

Overview

A pressing issue for helminth diagnostics is a reliable and affordable method to filter helminth eggs. Difficulty in filtration is attributed to the small size of helminth eggs; C. elegans specifically being about 50 micrometers. Currently, scientists use methods like the Kato Katz smear for diagnosis. This process involves taking a fecal sample, spreading it onto a microscope slide and examining egg morphology and characteristics. Additionally, traditional filtration methods may allow extra debris that could cloud the analysis of the sample or make the extraction process more tedious. This method is highly inefficient, expensive, subject to human error, and can take up to ten days to get results for. To solve this problem, Lambert iGEM decided to create a low cost and modular filter: MicroMesh. MicroMesh isolates helminths eggs from fecal and soil samples allowing users to run diagnostic tests in a more concentrated sample.

The workflow above describes the steps of a generic Kato-katz procedure.

Since isolation of helminth eggs precedes diagnosis, Lambert iGEM created MicroMesh, a modular feces filter that isolates eggs of any helminth species. According to the Center for Disease Control and Prevention (CDC), the eggs of soil-transmitted helminths are passed through the feces of infected people.

Lambert iGEM’s MicroMesh is based on a simple sieve; an object larger than the holes of the sieve will be trapped on top of the mesh. Based on this principle, the eggs are seperated from smaller microorganisms and larger debris. Using 3D printing, our product is efficient, easy-to-use, modular, and costs less than $2.

Design

Previous

We initially designed a straight cylinder model that would have two meshes to trap the specific sizes of particles in the middle. However, the design was not modular and inconvenient to use since it was a straight cylinder.

This shows a model of straight cylinder filter design.

This led us to modify this design to be smaller while still keeping the same fundamental principles. This would minimize the amount of filament used when printing while keeping the filter lighter and more portable. Additionally, we wanted to have the filter fit a microcentrifuge tube. With these ideas in mind, we redesigned the MicroMesh to be, in total, 75 mm tall, with the meshes having a diameter of 15 mm. Though this did make the MicroMesh significantly lighter and smaller, problems arose from it almost immediately. For one, we still had not addressed the problem regarding modularity. Secondly, the opening for the sample to go through was too small, meaning it got clogged up too easily and nothing would pass through while trials were being done. After testing, we realized that we did not need to have a filter the size of a microcentrifuge tube because we could just centrifuge down the sample after using the filter to remove the excess water. This led to the team concluding that though a smaller filter would be more convenient, the MicroMesh would have to be made bigger in order for the pores on the mesh to not get clogged by the sample.




This image shows two components of small funnel filter.

This image shows a small funnel filter assembled over a beaker.


Final Design

After seeing the results from the miniature funnel filter, we went back to a larger cylindrical design. We kept the funnel on the top. This time, we added a top sieve layer to help trap large soil particles, which was something that we suspected was clogging our mesh pores. This way, larger debris would be cleared before it even got to our first mesh, making the sample flow through easier. In addition, we addressed the idea of modularity. We decided to make disks for the mesh so they could be slid onto the pegs that were in place to keep the MicroMesh together. This way, the disks could be removed at any time. The mesh would have to be inserted inside of the disk for the mechanism to work.

We had to make the mesh disk this way in order to avoid gaps in the filter when it was assembled because putting the mesh directly onto the disk would cause a small gap the height of the mesh, which would cause the filter to leak. If leaking occurred, it could affect the concentration of helminth eggs in our sample. The addition of the disks also allows for the interchangeability of the mesh sizes. In the previous designs, a new filter would have to be printed every time the meshes were to be changed. With this update, the same filter could be used with several different disks. These new changes have helped evolve MicroMesh in order to achieve the design Lambert iGEM uses today. Our final design displayed below was optimized from a series of trial and error experiments of previous designs.




This image shows the MicroMesh's final design.

This image shows a small funnel filter assembled over a beaker.

This image shows the MicroMesh design schematic.


The MicroMesh is created with 3D printed thermoplastics, making it extremely cost-effective. The design of MicroMesh has one top-funnel component, one central cylinder and a bottom piece that attaches onto the cylinder to hold the mesh pieces in place. To keep the filter modular, we also designed a disk specifically for the mesh to be in. These pieces have mesh embedded inside a small disk that can be slipped onto the pegs of the cylinder. In addition, we designed the MicroMesh to have an optimized size where it is large enough for the sample to pass through while saving as much filament as possible.

The disk itself has a diameter of 65 mm and a total height of 5 mm. Since the mesh disks are not attached to the body of the MicroMesh, it allows for the disk to be able to be changed out based on what size mesh is needed. We inserted the mesh inside of the 3-D printed disk by sandwiching the mesh between two disks and gluing it together. For our MicroMesh specifically, we used meshes that were sized 75 μm and 44 μm to fit the range of C. elegans eggs that we were working with. The top component is a funnel in which a typical kitchen strainer was placed on top to separate out the larger debris found in soil. The end of the funnel includes pegs for the mesh disks to be slid on. The dimensions of the funnel component include the top opening, which has a diameter of 120 mm, the bottom opening, which has a diameter of 65 mm, and the pegs, which had a height of 12 mm and a diameter of 2.5 mm. The central cylinder is a basic cylinder that has four holes on the top and bottom of the cylinder to match the pegs on the top and bottom components. The cylinder has a diameter of 65 mm and 40 mm tall. Lastly, the bottom component is a simple disk similar to the disk for the mesh but has pegs instead of holes.

Theory

The theory behind the design of the MicroMesh is fundamentally simple. Since helminth eggs have very specific sizes, by creating a device that uses specifically sized mesh ranges, the eggs would be able to be trapped in the middle to move on with the rest of our workflow. A typical kitchen sieve is used first to separate out the larger debris found in the feces This will help prevent clogging of the smaller meshes. Then, the middle mesh layer of MicroMesh will trap any organisms or particles that are larger than 75 μm. This leads to the area in between two meshes: the middle and bottom one. This layer will contain particles between the sizes 44 μm and 75 μm, which is a size range that fits C. elegans eggs. Lastly, the third layer of mesh will let any matter smaller than 44 μm to pass through, including the water being passed through MicroMesh. This allows the remaining sample trapped in the middle of the MicroMesh to be within the range of 44 μm and 75 μm, which will contain helminth eggs to move on with the rest of our workflow.

Assembly

Click here to access the STL files.

  1. Push the larger micron mesh filter disk up onto the four pegs that are attached to the funnel
  2. Insert the middle cylindrical component onto the funnel pegs so that it fits snugly
  3. Push the smaller micron mesh disk down onto the thin bottom disk with pegs
  4. Insert this piece into the other end of the cylindrical middle component

This is the tutorial of MicroMesh construction and use.

Protocol

Optimized Fecal Protocol

  1. Assemble the MicroMesh appropriately: (The funnel, the larger sized micron mesh disk, the middle component, the smaller micron disk, then the disk with solely pegs)
  2. Measure 2g of sample and mix with 5 mL of water**
  3. Vortex sample until the sample is homogeneous or create a homogenous solution by using a mortar and pestle**
  4. Pour sample over the filter
  5. Pour 10 mL of water over sample***
  6. Wait until all of the sample is filtered through the filter and there is no more remaining water above the larger micron filter
  7. Disassemble the filter and take the smaller micron filter disk
  8. Flip disk over and pour 12 mL water over sample above a beaker
  9. Take sample and centrifuge down and remove the excess water on the top
  10. The sample is now prepared for the bead homogenizer

**varies per sample due to materials in sample and consistency,
***pour more water until all remnants of the sample that can pass through the uppermost layer pass through

It is important to note that when making a homogenous solution, the amount of water necessary to achieve a resulting solution does not have to remain constant between samples to ensure that helminth eggs are trapped on the smaller filter. For more difficult samples we suggest running more water over the filter to lift any particle blocking, for example, the larger of the two-micron filters, to allow for the rest of the sample to flow through.

Results

Testing at Barr and Ryan Lab at Emory

Our lab at Lambert IGEM is biosafety level 1, meaning we are not able to deal with fecal matter. We visited the Barr and Ryan Lab at Emory University in September of 2019 to test fecal samples through the MicroMesh. While at the lab, we tested MicroMesh on feces from Macaca fascicularis and solid-eating infant feces. This experience allowed us to confirm our theory that MircroMesh would be able to filter through fecal samples and also create an optimized protocol outlined above.

In order to analyze results, we had to confirm that the particles trapped between the two mesh layers were within our mesh ranges, we looked at the particles through a microscope and measured the sizes. To do this, we took the samples that we got from MicroMesh, which had water in them since we used water to get them off the small mesh, and we centrifuged them down to compact the solids. We removed the excess water and then made microscope slides with the remains of the sample. We took pictures of the slides under a microscope and then used a software to determine the sizes.

Lambert iGEM is testing the efficiency of MicroMesh using Macaca fascicularis monkey feces.

This is a picture of Lambert iGEM at Barr and Ryan Lab at Emory University.

Data

Our proof of concept model organism, C. elegans, measures approximately 50 μm in length (Riddle et al.). To account for this measurement, our team used 75 μm-44 μm mesh to test. Due to our lack of ability to secure C. elegans eggs, we filtered 50 μm glass beads.

The data from fecal samples comes from the fecal samples we collected from the Ryan and Barr Labs when we were there. While we were there, we conducted four trials total: two for the Macaca fascicularis feces and two for the infant feces. The data we collected is the sizing of the particles trapped on the large and small mesh.

In order to collect data for this used the software Image J which allowed us to collect data on what the size of the particles in the sample based on the field of view. We used a 4x objective lens to analyze our samples. In order to calculate the field of view we multiplied the 10x eyepiece magnification by the 4x objective lens magnification and divided by 18, the field number of the microscope we used. The field of view we used was 450 micrometers

Small Filter Sample Type Length in Micrometers
cynomolgus feces 1 52.0
cynomolgus feces 2 69.3
cynomolgus feces 2 51.8
infant feces 1 72.9
infant feces 2 58.3
Large Filter Sample Type Length in Micrometers
cynomolgus feces 1 76.5
cynomolgus feces 1 78.4
cynomolgus feces 2 100.9
infant feces 1 99.0
infant feces 1 81.6
infant feces 2 88.4
infant feces 2 90.3

The 1 and 2 indicate which trial the data is from and the addition of data from more than one trial was due to the appearance of numerous particles on one slide.

In order to get representative data on if MicroMesh would efficiently work with helminth eggs, we obtained 50 micron glass beads, which is the ideal size for helminth eggs. Using the beads, we tested if the beads would get trapped on the small filter. To do this, we placed a small amount of the beads onto the top layer of the MicroMesh and poured water through it to simulate what we would do with remnants of fecal matter that lay on the top of the filter. Then we waited until all of the water flowed through MicroMesh and then disassembled it in order to check if the beads would be resting on the second micron mesh. We observed that the beads were caught on the second micron mesh; therefore we knew that MicroMesh would adequately trap helminth eggs of this size.

While observing the filter, we realized that when we were attempting to take a picture under the microscope, the 50 micron beads were not visible as they were white and lacked color. This meant that they could not be seen under the harsh light and mesh background. Therefore we decided to dye the beads and repeat the process of obtaining data in which we had the beads trapped along the small mesh.

This shows the top view of MicroMesh with 50 micron glass beads (dyed purple for visibility).


OPENCELL

Overview

After the filtration of helminth eggs, DNA must be extracted from the eggs in order to move forward in our LABYRINTH workflow. However, helminth eggs possess a rigid chitin middle layer that is notoriously difficult to break, making our process of extracting DNA all the more difficult. Chiefly, a bead homogenization device must break down the chitin layer, while preventing shearing or damage to the DNA. Currently, by quantifying DNA via a Nanodrop, we can confirm the presence of usable DNA and through a successful PCR product.

With our main goal being frugality, the OpenCellX system needs to employ unique mechanics for maximum efficiency. Over time, the design has slowly evolved to use less energy, and lyse samples with as much efficiency as possible. The end product must be a machine applicable to various workflows and in multiple labs, not just to our project, and show comparable results to commercial solutions.

Design

Previous

The very first homogenizer design was based off the motion of desktop vortexers, which use an imbalanced rotation to provide high-velocity oscillations. To reproduce a similar effect on a 3D Fuge, a small counter weight was added to the circumference of the device. After even basic prototyping it was clear that no reasonable amount of energy could be applied to the system to create strong enough oscillations or maintain a high RPM.

The Fan-Based homogenizer used a fan to create an oscillatory motion that we presumed would homogenize the samples.

From here, we concluded that a reproducible and stable system would be easiest to power from a small DC motor rather than by human power. Inspired by the USB fan based vortexing systems, we designed a scotch-yoke mechanism with a new computer fan system at its heart. As the fan rotates, an arm affixed to the fan hub would power the stroke of the mechanism, sliding an extremely thin plate back and forth along linear rails. While the machine was significantly more promising than our preliminary 3D Fuge concept, it still lacked the efficiency required for an on site or in lab device, unable to provide usable results after even three or four hour runs. Because of its strictly linear motion, much of the original sample would stick to the sides of the tube and by the end, we would have less than 50% of the matter homogenized. Furthermore, we analyzed the motion through a high-speed camera and discovered that the scotch-yoke stroke applied varying load on the fan over the course of one cycle. As a result, the fan was unable to maintain a high speed while also using a large amount of energy.

The planetary gearbox design was separated into three parts: the base, which contained the internal rotating mechanism, the supporting secondary layer that kept the rotating arm stable, and the gear lid that powered the homogenizer.

This version of OpenCell was sent to GSU as a collaboration for their project, and to provide data on ours.


We arrived at our new design with inspiration from the epicyclic gear-train found in many mechanisms. Our initial design involved a fixed central gear, two secondary idle gears, and finally, two driven tertiary gears to which the microcentrifuge tubes are attached to. When an arm is revolved around the central gear, the rotation of the idle gear contributes to a counteracting rotation on the driven gear, and as a result, the tertiary maintains a constant relative position throughout the entire cycle. Therefore, the centripetal force generated by the revolution of the tube forces beads around the entire internal surface of the tube due to their inertia. We analyzed the motion of our most successful design, finding that the beads reach their highest velocity in the tube at 4 key points which effectively homogenize the sample.

Final

OpenCell is created with 3D printed thermo-plastics, making it very inexpensive to produce. The final OpenCell design features a wide circular base with a radius of 75.6 mm and a fixed central gear raised above the base with a rectangular slot for a locking key. On the circumference, there is a cylindrical extension where the motor of diameter 2.6inch can be inserted facing down. The motor turns a 15 tooth gear affixed directly to the motor shaft, which consequently drives the main 45 tooth gear which revolves around the central gear.


This image shows the functional device used for testing.




Advantages

By using 3D printed technology, we are able to offer OpenCell at a fraction of the price of industrial homogenizers, making our device more practical for underfunded research labs diagnostic centers. Applying basic lubricant to the surface of our homogenizer will reduce both friction between the pieces, increasing the RPM and efficacy of the device, but also prevent immediate wear and tear of the components. Capable of running on batteries, OpenCell is the first homogenizer that can operate without the crutch of 120V wall socket electricity.

Within the 45 tooth gear, the main planetary gearing revolves internally around the central gear. The secondary idle-gears are fitted on either side of the central gear, and the tertiary gears and fitted on the other side of the secondary idle-gears. The supporting arm is fixed atop of the gears, acting as a stabilizing structure for the system, while also separating the tube holders, which are fixed onto the tertiary gears. Finally, a lid encloses the entire mechanism protecting both the user from the high speed parts as well as the mechanism from dirt and debris.

Simply put, the cost of most lab equipment is just too high for many labs to be able to use. Especially in underdeveloped areas like the Dominican Republic, that our team has had the opportunity to visit, who require them to diagnose and treat illnesses. Of all commercial lab instruments, centrifuges, vortex mixers, and homogenizes are some of the most essential, yet each one may cost upwards of $500, or upwards of $1000 for a bead-beater. The goal of OpenCell is to combine the utility of all three machines into one, frugal system.

Using a powerful, yet low-cost dc motor, the OpenCell Bead-Homogenizer operates at upwards of 1200 RPM to expose the DNA of many specimen, then the same mechanism can be used without beads as a vortex mixer. To reach speeds high enough for proper centrifugal, the OpenCell employs a removable motor enclosure to which a centrifuging attachment can be affixed. Because of this, many lab workflows and procedures can be run with the OpenCell at the heart of the process, and all at just a fraction of the cost. The frugal OpenCell device is targeted towards underfunded labs and on-field applications, and will cost around $5, less than 1% of the price of desktop bead homogenizes. Its small motor can be run efficiently on off the shelf 9v batteries, or with a basic 9v ac adapter (the same kind used for lab balances). Yet, regardless of the low cost, our results have shown to be comparable to conventional bead-beaters.

Another area in which we could save money is with reagents and supplies. DNA extraction kits from companies like Qiagen and Bio-Rad can cost upwards of $5 per sample, the same price as the OpenCell itself. To reduce cost, the OpenCellX system employs a protocol using standard eppendorf tubes and low cost steel beads to bring the cost per sample down to just 50%. Moreover, incorporating non-proprietary lysis buffers and reagents, that cost can be brought down further.

Our results have shown that low cost steel beads and standard tubes provide equal, though usually better, DNA yields for a fraction of the price, using OpenCell. Furthermore, trials run on the bead beater provided to us by the GSU iGEM team had shown a similar trend.

Assembly

Click here to access the STL files.

  1. Ensure that all parts are available and separate: 1 45 tooth gear, 2 10 tooth gears (with holes), 2 10 tooth gears (with protruding pegs), 1 rotating arm, 1 central stabilizing key, 2 test tube holders, 1 motor/battery mount, 1 15 tooth gear, 1 motor locking key
  2. Fit the larger 45 tooth gear onto the base of the homogenizer, making sure that the hole in the center of the gear fits around the raised fixed base gear.
  3. Fit the secondary gears (smaller 10 tooth gears with holes in their centers) around the protruding pegs located on either side of the fixed 15 tooth base gear.
  4. Fit the tertiary gears (small 10 tooth gears with protruding pegs in their centers) next to the secondary gears. As a result, each secondary gear should have 1 tertiary gear on one side and the base gear on its other side.
  5. Fit the rotating arm (12.5 cm rectangular beam) onto the gears so that the holes on each end of the arm can be fitted onto the pegs on two ends of the main gear and so the internal holes can be fitted onto the pegs of the secondary gears.
  6. Lock the mechanism by inserting the stabilizing key into the slot in the middle of the stationary base gear.
  7. Insert the two test tube holders atop the tertiary gears.
  8. Fit the motor (axle pointing down) into the circular motor shaft and fit the battery into the rectangular shaft. Ensure that the motor and battery are wired.
  9. Slide the motor/battery mount into place on the extended rectangular base of the homogenizer base, making sure that the slot for the key on the mount lines up with the slot on the rectangular base.
  10. Insert the motor locking key into the key slot to lock the mount into place.
Protocol

DNA Extraction from Helminth Eggs

The Following Protocol is used for DNA extraction from helminth eggs:

  1. Notes before starting
    • Perform all centrifugation steps at room temperature (15–25°C).
    • If Solution C1 has precipitated, heat at 60°C until precipitate dissolves.
  2. Add 0.25 g of soil sample to the PowerBead Tube provided. Gently vortex to mix.
  3. Add 60 μl of Solution C1 and invert several times or vortex briefly.
    • Note: Solution C1 may be added to the PowerBead tube before adding soil sample
  4. Secure PowerBead Tubes horizontally using a Vortex Adapter tube holder (cat. no. 13000–V1–24).
  5. Vortex at maximum speed for 10 min.
    • Note: If using the 24-place Vortex Adapter for more than 12 preps, increase the vortex time by 5–10 min.
  6. Centrifuge tubes at 10,000 x g for 30 s.
  7. Transfer the supernatant to a clean 2 ml collection tube. Note: Expect between 400–500 μl of supernatant. Supernatant may still contain some soil particles.
  8. Add 250 μl of Solution C2 and vortex for 5 s. Incubate at 2–8°C for 5 min.
    • Note: You can skip the 5 min incubation. However, if you have already validated the DNeasy PowerSoil extractions with this incubation we recommend you retain the step.
  9. Centrifuge the tubes for 1 min at 10,000 x g.
  10. Avoiding the pellet, transfer up to 600 μl of supernatant to a clean 2 ml collection tube.
  11. Add 200 μl of Solution C3 and vortex briefly. Incubate at 2–8°C for 5 min.
    • Note: You can skip the 5 min incubation. However, if you have already validated the PowerSoil extractions with this incubation we recommend you retain the step.
  12. Centrifuge the tubes for 1 min at 10,000 x g.
  13. Avoiding the pellet, transfer up to 750 μl of supernatant to a clean 2 ml collection tube.
  14. Shake to mix Solution C4 and add 1200 μl to the supernatant. Vortex for 5 s.
  15. Load 675 μl onto an MB Spin Column and centrifuge at 10,000 x g for 1 min. Discard flow through.
  16. Repeat step 14 twice, until all of the sample has been processed.
  17. Add 500 μl of Solution C5. Centrifuge for 30 s at 10,000 x g.
  18. Discard the flow through. Centrifuge again for 1 min at 10,000 x g.
  19. Carefully place the MB Spin Column into a clean 2 ml collection tube. Avoid splashing any Solution C5 onto the column.
  20. Add 100 μl of Solution C6 to the center of the white filter membrane. Alternatively, you can use sterile DNA-Free PCR Grade Water for this step (cat. no. 17000–10).
  21. Centrifuge at room temperature for 30 s at 10,000 x g. Discard the MB Spin Column. The DNA is now ready for downstream applications.
    • Note: Solution C6 is 10 mM Tris-HCl, pH 8.5. We recommend storing DNA frozen (–20° to –80°C) as Solution C6 does not contain EDTA. To concentrate DNA see the Hints & Troubleshooting Guide.

Adjustments to Qiagen's DNeasy PowerSoil Kit

The protocol of Qiagen’s DNeasy PowerSoil Kit was utilized for the extraction of C. elegans egg DNA with several adjustments:

  1. Add 250 uL of culture to power bead tube then add 60 uL of solution C1 and freeze for 35 minutes at -80°C.
  2. Heat in 70°C water for 5 minutes and homogenize for 15 minutes.

All centrifugation steps are at 12,000 x g rather than 10,000 x g, and all centrifugation lengths prior to the addition of solution C4 are doubled.


DNA Extraction from Spinach Leaves

The following protocol is used for DNA extraction from spinach leaves:

  1. Add 5–100 mg of plant tissue and 500 µl Solution CD1 to a 2 ml tissue disruption tube. Vortex briefly to mix.
    • Note: We recommend that the tissue be cut into small pieces before loading into the bead tube. For tough plants or seeds, pregrinding the material with a mortar and pestle may increase yield.
    • Note: If your sample is high in phenolic compounds, add 450 µl CD1 and 50 µl Solution PS. For problematic samples, you can add up to 100 µl Solution PS and correspondingly decrease CD1 at this step.
  2. Homogenize using the OpenCell homogenizer. Note: Homogenization speed and duration may need to be optimized for your specific sample to ensure highest DNA yield and quality.
  3. Centrifuge the Tissue Disruption Tubes at 12,000 x g for 2 min.
  4. Transfer the supernatant to a 1.5 ml collection tube (provided). Note: Expect 350–450 µl. The supernatant may still contain some plant particles.
  5. Add 200 µl Solution CD2 and vortex for 5 s. Note: For problematic samples, you can add up to 250 µl Solution CD2 at this step.
  6. Centrifuge at 12,000 x g for 1 min at room temperature. Avoiding the pellet, transfer the supernatant to a 1.5 ml collection tube (provided). Note: Expect 400–500 µl.
  7. Add 500 µl Buffer APP and vortex for 5 s
  8. Load 600 µl of the lysate onto an MB Spin Column and centrifuge at 12,000 x g for1 min
  9. Discard the flow-through and repeat step 8 to ensure that all lysate has passed through the MB Spin Column
  10. Carefully place the MB Spin Column into a clean 2 ml collection tube (provided). Avoid splashing any flow-through onto the MB Spin Column.
  11. Add 650 µl Buffer AW1 to the MB Spin Column. Centrifuge at 12,000 x g for 1 min.
  12. Discard the flow-through and place the MB Spin Column back into the same 2 ml collection tube.
  13. Add 650 µl Buffer AW2 to the MB Spin Column. Centrifuge at 12,000 x g for 1 min.
  14. Discard the flow-through and place the MB Spin Column into the same 2 ml collection tube.
  15. Centrifuge at up to 16,000 x g for 2 min. Carefully place the MB Spin Column into a new 1.5 ml collection tube (provided).Add 50–100 µl of Buffer EB to the center of the white filter membrane.
  16. Centrifuge at 12,000 x g for 1 min. Discard the MB Spin Column. The DNA is now ready for downstream applications.
    • Note: We recommend storing the DNA frozen (–80 to –20°C), because Buffer EB does not contain EDTA.
    • Note: Buffer EB is 10 mM Tris (pH 8.0). The DNA bound to the MB Spin Column membrane is solubilized into Buffer EB.

Adjustments to Qiagen's DNeasy Plant Pro Kit

The protocol of Qiagen’s DNeasy Plant Pro Kit was utilized for the extraction of spinach leaf tissue DNA with a few adjustments:

  1. 50 mg of plant tissue, 600 µl of solution CD1, and 8 3.2 mm chrome-steel beads are added to a microcentrifuge tube. This standardizes the protocol for spinach leaves.
  2. The OpenCell homogenizer homogenizes the sample for 6 minutes.
Results

Spinach





Processing DNA samples using the Qiagen protocol, reagents, and supplies served as a baseline test from which we could improve our own protocol. Samples were each run for 2 minutes and were quantified using a nanodrop. Although the mean yield from OpenCell was slightly lower, the results are within the margin of error calculated through SEM. The DNA purity results, on the other hand, favored the OpenCell but again all results are within the margin of error.






To tune the OpenCell protocol, we began with testing the time required to properly lyse a spinach sample. Running sample triplicates at 1, 3, 5, 7, 9, and 11 minutes, we tested the Yield and Purity of DNA using a nanodrop. According to the data, there is a positive trend, but after the 5-minute mark there is no significant difference between samples. Therefore, 5 minutes is the most efficient time to provide a significant DNA Yield for further use. For purity, there is no significant difference between any of the times, however there is a trend for more consistent results as the time is increased.




Using the optimal protocol for each machine, the mean Spinach DNA yield from the OpenCell design was slightly higher than that of the BioSpec Mini-BeadBeater, though it is still within the margin of error. This is likely due to the small number of trials run on either homogenizer. The purity of either machine shows a similar trend, however the data is much more consistent and therefore has a smaller error. We plan to run more trials, and further tune our protocol to provide better, and more consistent results.

Algae


Georgia State University's iGEM team tested our frugal bead homogenizer and compared results with their commercial homogenizer. They gave us significant feedback on ease of use and functionality and provided insight into potential improvements to our design. After testing, they described our frugal homogenizer results as "better than the commercial homogenizer" in terms of their experiment, resulting in greater cell viability. Our homogenizer was less aggressive than the commercial one with more tuneability and versatility, yielding live and motile cells.


GSU homogenized microalgae with a BioSpec Mini-BeadBeater. No live cells are present.

GSU homogenized microalgae with OpenCell. Live and motile cells are present.


Helminths


Homogenization Method Yield (μg) Purity (A260/A280)
Vortex Mixer-Qiagen Garnets 0.135 1.37
OpenCell-Zirconium Beads 100 Micron 0.276 1.62
OpenCell-Qiagen Garnets 0.69 1.77
OpenCell-Qiagen Garnets after PCR Purification 1.560 1.74

The DNA extraction from C. elegans eggs was significantly better after the adjustments to the protocol as well as extending the time of homogenization. As shown above in the results, Opencell worked much better in breaking through the chitin layer in comparison to the regular vortexer.


After extraction of DNA from the C. elegans, it was amplified by PCR. The entire genomic DNA of the C. elegans was extracted in the sample, but the PCR was only done with the forward and reverse primers of the trigger sequence to the toehold. A gel was run to confirm the PCR. The gel proved that the amplification of the trigger sequence was successful. After the PCR, a PCR purification was done using the Monarch® PCR & DNA Cleanup Kit (5 μg) in order to purify the amplified DNA.

Gel with amplified DNA from PCR in lane 4 and a 100bp ladder in lane 8. The band around the 100 base pair band on the ladder indicates a successful PCR

After the PCR, protocol for a PCR cleanup was done to purify the amplified DNA. Following the amplification of the DNA, the next step would be to use the HiScribe T7 High Yield RNA Synthesis Kit to transcribe the trigger DNA into RNA in vitro with the use of T7 RNA Polymerase. Lambert iGEM plans to do this step in the future. After converting the resulting DNA to RNA, the single stranded RNA trigger sequence would be introduced to the toehold created by Lambert iGEM and bind to the toehold, causing the hairpin loop of the toehold to unravel and allow for translation and expression of the reporter gene GFP, ultimately proving the device's effectiveness on even the most resiliant organisms.


OPENCELL PRO

Overview

In a proper lab situation, fine control of RPM and other parameters is paramount in repeatability of workflows and the success of the experiment as a whole. While our frugal, in field, machine was designed for portability and accessibility, the OpenCell Pro can serve any underfunded lab with features to ensure consistent results and easier use.

OpenCell Pro targets a different segment of consumers, and as such is more expensive than the base model. However, the increased costs produces an overall experience more akin to currently available homogenization equipment. With more premium features, OpenCell Pro is offered as a pre-built device that resembles typical commercial homogenizers.

Design

OpenCell Pro contains many components not present in the frugal OpenCell design. A larger motor will accelerate workflows, and improve the machines utility in centrifugation applications by increasing the load speed 10,000 RPM. Powered by a proper motor driver, the user will be able to set accurate speed targets to improve consistency. On the exterior of the device is a 2 row LCD display with an intuitive dial and button navigation system. At the heart of the electronic system is an Arduino microcontroller which runs PID voltage control, enables time based runs, and can even keep track of trials.

While the new electronics make the system simpler to use, the hardware which the system employs stays the same and the efficient homogenization results from the basic model are only going to be improved in the Open Cell Pro.

Utilizing an Arduino Uno and a seperate motor driver, the OpenCell Pro can automatically run for set times and adjust speeds for a tuneable operation and broader application.

This advanced OpenCell Pro design will process four tubes, instead of two, and replicates the interface of conventional lab instruments.

REFERENCES

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[9] Patil, M. D., Patel, G., Surywanshi, B., Shaikh, N., Garg, P., Chisti, Y., & Banerjee, U. C. (2016). Disruption of Pseudomonas putida by high pressure homogenization: a comparison of the predictive capacity of three process models for the efficient release of arginine deiminase. AMB Express, 6(1). doi: 10.1186/s13568-016-0260-6

[10] Periago, M. V., Diniz, R. C., Pinto, S. A., Yakovleva, A., Correa-Oliveira, R., Diemert, D. J., & Bethony, J. M. (2015). The Right Tool for the Job: Detection of Soil-Transmitted Helminths in Areas Co-endemic for Other Helminths. PLOS Neglected Tropical Diseases, 9(8). doi: 10.1371/journal.pntd.0003967

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