Mar 11, 2026
Zoospore microfluidic chemotaxis assay
- Eric McLamore1,
- Diana Vanegas1
- 1Biological and Agricultural Engineering, University of Arkansas
- SNAPS research group
Protocol Citation: Eric McLamore, Diana Vanegas 2026. Zoospore microfluidic chemotaxis assay. protocols.io https://dx.doi.org/10.17504/protocols.io.5jyl84297g2w/v1
Manuscript citation:
Protocol M1.1: Microfluidic soil-pore chemotaxis demonstration with plant roots
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: March 09, 2026
Last Modified: March 11, 2026
Protocol Integer ID: 308276
Keywords: Phytophthora, microfluidic, infection, plant , root, water mold, pathogen, chemotaxis of motile phytophthora zoospore, microfluidic chemotaxi, motile phytophthora zoospore, microfluidic device, chemotaxi, cost microfluidic device, plant root, filled soil
Funders Acknowledgements:
National Institutes of Food and Agricultural Sciencees Multistate
Grant ID: NC1194
Abstract
This protocol demonstrates chemotaxis of motile Phytophthora zoospores toward plant roots using a low-cost microfluidic device that mimics water-filled soil pores.
Guidelines
* The experiment requires simultaneous germination of seeds and preparation of motile zoospore suspensions from cultures Phytophthora.
Careful planning must be followed to ensure timing of each prep.
Experiments should be well-organized and planned at least 7-10 days prior to the event
Materials
Biomaterials
- Zoospore suspension (prepared from Phytophthora capsici Leonian 52771 using zoospore preparation protocol)
- Cucumber seeds from reliable source (local organic market or nursery).
Note
Alternatively, pepper seeds may be used. Pepper is one of the best-studied hosts for P. capsici. Pepper seeds tend to germinate slower and less with uniformity. Germination is typically improved by using a heating mat but still not as high of a rate when compared to cucumber.
Chemicals
- 70% (v/v) ethanol
- 20% (v/v) bleach
- Autoclave-sterilized water or DI water
- Calcium chloride solution (1.0 mM CaCl₂), approximately 40 mL for one group is sufficient.
Hardware
- Cameras or video recording devices (at least 10-30 fps is suggested)
- Phase-contrast compound microscope or other capable instrument (zoospores are 8-12 µm in diameter). See table below for magnification recommendations
- Micropipettes (or transfer pipettes)
Software
- imaging software for camera/microscope (if used)
Troubleshooting
Safety warnings
Safety information
Personal Protective Equipment (PPE)
- Wear lab coat at all times
-Wear gloves at all times
-Wear certified laboratory eye protection at all times
-Avoid use of phone or other personal items during lab (e.g., keys, writing pens, etc.)
Safety information
Handling biomaterials
- Handle plant pathogens using standard plant pathology laboratory practices.
- Avoid environmental release of cultures or infected plant material.
- Avoid contact with microbial cultures or staining reagents.
Safety information
General safety
- Conduct work involving staining dyes or solvents in a well-ventilated area.
- Avoid open flames near solvents or dyes.
- Dispose of biological materials according to institutional biosafety procedures.
Before start
This protocol requires use of Zoospore preparation protocol prior to the lab. This preparation requires approximately 2-3 days prior to the protocol herein. The microfluidic device consists of a narrow channel opening into a wider root-tip observation zone. Students should focus on aggregation behavior of zoospores close to the root tip rather than only on movement through the channel.
(optional) If fabricating custom microfluidic channels in glass slides, use our protocol based on a benchtop UV laser engraver.
PREPARATION
(optional) Prepare microfluidic glass slides using UV laser engraver
Follow the Zoospore preparation protocol
-Carefully follow the zoospore preparation protocol to obtain motile Phytophthora capsici L suspensions from culture.
-Plan experiments carefully to ensure timing of cultures for lab experiment
Figure 1. Sporangium of Phytophthora capsici releasing zoospores. The Oval sac is called the sporangium. The narrow tube is the discharge papilla and the small black dots exiting the structure near the base of the papilla are the zoospores.
Sterilize seeds
- Soak dry seeds in 70% (v/v) ethanol for 1 min
- Remove seeds and sterilize in 20% (v/v) bleach for 10 min
- Immediately wash seeds in DI water (or sterilized water) by agitating gently for 1 min
- Pour off the DI water, and repeat the washing process for at least 1 min
Place seeds in germination chamber
- Soak a KimWipe or clean paper towel in 1.0 mM CaCl₂
- Place sterile seeds in a dilute calcium chloride solution (0.5 to 1.0 mM CaCl₂)
Note
Use of 1.0 mM CaCl₂ (as opposed to DI water or other buffer) is important for this step to ensure:
- stable root cell membranes
- 1.0 mM CaCl₂ helps maintain gentle ionic conditions compatible with root viability and zoospore chemotaxis assays.
- protocol is compatible with classic zoospore chemotaxis assays
- Germinate sterile seeds for 24–36 h in until roots reach 1–2 cm.
Figure 2. Clean, unused plastic food containers (or Ziploc bags) are good germination chambers for small batches of seeds.
Note
For large labs, use a sterilized seed sprouter tray to germinate dozens/hundreds of seeds simultaneously.
If using this approach, it is recommended to germinate two batches as a precaution (redundancy in case of contamination to a large batch)
Store germinating seeds
-For best results, store the cucumber seedlings in the refrigerator for up to 48 hours prior to use. This will slow down the rapid development phase and ensure the roots are at the appropriate growth stage for the experiment
-See the Table below for appropriate storage times of germinating seeds, and plan the experiments carefully
| Storage condition | Usable time window | |
| room temperature (24–26 °C) | 12–24 hours | |
| cool room (15–18 °C) | 24–36 hours | |
| refrigerator (4–8 °C) | up to ~48 hours |
Table 1. Practical storage window for 48-hour cucumber seedlings.
Prepare microfluidic device and imaging system
Arrange microfluidic devices
- Collect two microfluidic slides per group (either the commercial devices or the laser-engraved custom devices)
- Depending on the specific design of the microfluidic slide, multiple wells may be included for control studies (e.g., latex beads as non-target particle, DI blank, challenging system with other chemicals, etc.)
FIgure 3. The ibidi microfluidic sytem is an excellent option for smaller groups
Prepare cameras or video recording devices for each station
- Prepare microscopes with at least a 20X magnification. See Table 2 for magnification requirements
| Objective | Use | |
| 10× | find root / field | |
| 20× | best overall view of chemotaxis | |
| 40× | detailed zoospore observation |
Table 2. Magnification requirements for microscope used in zoospore microfluidic experiments
Note
Depending on the learning objective, various microscope options are available
depending on goal of lab)
- A phase-contrast compound microscope with at least 20X magnification is preferred for this lab.
- If not available, a Differential Interference Contrast (DIC) or alternatively a Brightfield microscope with reduced illumination is sufficient.
- A low-cost option is to use a USB camera (link). However, the images will be limited to clustering of zoospores at root tip (small black dots, grainy images).
Prepare buffer
- Ensure each team has approximately 5mL of 1.0 mM CaCl₂ for experiments
- Ensure access to pipettes at each station
Arrange seedling with microfluidic device
Assemble the microfluidic device on a clean platform
- Place the microfluidic device on the acrylic plate
- This important step ensures that the device may be easily re-located at later stages of the protocol
Figure 4. Place microfluidic device on clean acrylic sheet for easy re-location
Carefully fix seedling to microfluidic slide
- Place seedling onto microfluidic slide
- Carefully position the root tip near the outlet side of the microfluidic slide
- Use biocompatible adhesive (e.g., guar gum) to fix seedling
Figure 5. Lay seedling flat on surface os that root tip is aligned with center of well. Use guar gum or other soft polymer to gently fix the root to the glass slide. If using a microfluidic design as shown in the diagram, the two inlet wells may be used for replicates, blanks, or other controls.
Note
When fixing the root to the glass slide, be gentle. Damage to the root can change the chemistry at the tip and alter the experiment
Fill inlet well(s) with electrolyte
- Using a pipette, fill inlet wells of microfluidic device with 1.0 mM CaCl₂
- Be mindful and avoid injecting any bubbles into the microfluidic
Figure 6. Fill each well of the microfluidic device with with 1.0 mM CaCl₂. Inset shows root tip submersed in electrolyte solution
- As the electrolyte solution flows from the inlet to the root observation zone (outlet), be careful not to damage the root tip
- After filling well, gently position the root tip so it is submerged in the solution
Note
Important step: After the root tip is submerged, allow the device to sit undisturbed for 5–10 min before adding zoospores.
Perform final check of assembly
Perform a final check of the setup prior to beginning the next section
Note
1) Is the device stable on the counter?
-If not stable, level the acrylic plate prior to proceeding
2) Is the guar gum adhering well, and keeping the root in position?
-If adhesive is not functioning well, reposition carefully and avoid damage to root
3) Is the root tip submersed in the root observation zone (outlet well)?
-If tip is not submersed, use plastic-tipped tweezers to gently re-position without squeezing
4) Is the root tip near the center of the root observation zone and not adhering to the wall?
-If tip is not centered, use plastic-tipped tweezers to gently re-position without squeezing
5) Is the root tip damaged?
-note damage in lab notebook. If possible, replace for non-damaged seedling
BEGIN EXPERIMENT WITH ZOOSPORES
Add zoospore suspension
BEGIN EXPERIMENT WITH ZOOSPORES
Gather zoospore suspension
- Collect zoospore suspension
- Gently invert 2-3 times to mix suspension
Note
Use only freshly prepared suspensions showing active motility under a microscope before distribution to students.
BEGIN EXPERIMENT WITH ZOOSPORES
Add zoospores to microfluidic device
- Carefully pipette 5µL of zoospore suspension into the inlet well of the microfluidic
- Inspect the root tip to be sure it has not moved
- If using a microfluidic with multiple inlet ports, experiments may be conducted for challenging with system with other compounds/particles/cells
Figure 7. Carefully pipette 5µL of zoospore suspension into the inlet well of the microfluidic.
Note
The device consists of a narrow microfluidic channel opening into a wider observation zone where the root tip is located. Students should focus on aggregation behavior close to the root tip rather than only on movement through the channel.
Additionally, students should observe that fluid flow is not forcing the zoospores through the device, the movement is due to chemotaxis
OBSERVATION
Record observations
Observe movement of zoospores toward root observation zone (outlet)
- Using a 10X objective, observe zoospore movement through channels.
Note
- Start at 10× to locate root and channel
- Move to 20× for tracking approach behavior
- Move to 40× only after spores are visible near the tip
- The zoospores are much smaller than the channel width (drawing not to scale)
Note
General sizes for reference:
Zoospores = 8-12 µm
Channel width = 100 µm
Learners may observe small dark specks moving through the channel toward the root region. Near the root tip, movement may slow, turning may increase, and local accumulation may occur.
Figure 8. Observe zoospores moving through the microfluidic channel. Drawing is not to scale.
Note
Observation checkpoint 1: channel
- Are zoospores visible?
- Are they motile?
- Are they moving randomly, drifting, or orienting toward the root region?
Observe zoospores near root tip
- After observing the microfluidic channel for approximately 5 min, reposition the microscope to inspect the root tip
- Observe root tip under microscope
- Record aggregation behavior near root tip.
Figure 9. Observe zoospores aggregating near root tip. Drawing is not to scale.
Note
Observation checkpoint 2: near-root region
- Is there visible accumulation near the tip?
- Is accumulation strongest at the very tip or slightly behind it?
- Is motion slower or more clustered near the root than in the channel?
CLEANUP
Dispose of biological materials appropriately
- Flush microfluidic system with the cleaning solution provided by instructor
- Inspect device under microscope to ensure channels are clean and do not contain residue or zoospores
- Clean workstation surfaces and acrylic plate with laboratory cleaning solution provided by Department
- Ensure all pipette tips are disposed according to lab standard operating procedures (SOP)
- Store electrolytes according to instructor workflow
- Dispose of all containers in the BIohazard according to local SOP
- Turn off microscopes and electronic equipment.
DATA MANAGEMENT
Properly name files and store in shared folder
- If images are captured, save and name the file using the following: "ZOOSPORE_MICROFLUIDIC_date."
- Store images and videos from experiments in laboratory shared folder assigned by instructor