Jun 29, 2026

High-throughput Larval Development Assay (HTLDA)

 Forked from High-throughput Larval Development Assay (HTLDA)
DOI
https://dx.doi.org/10.17504/protocols.io.5qpvoejydl4o/v1
  • 1Johns Hopkins University
Erik Andersen
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DOI: https://dx.doi.org/10.17504/protocols.io.5qpvoejydl4o/v1
Protocol Citation: Amanda Shaver, Erik Andersen 2026. High-throughput Larval Development Assay (HTLDA). protocols.io https://dx.doi.org/10.17504/protocols.io.5qpvoejydl4o/v1
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
This protocol is active in the lab.
Created: May 19, 2026
Last Modified: June 29, 2026
Protocol  Integer ID: 317464
Keywords: C. elegans, high-throughput larval development assay , C. briggsae, C. tropicalis, throughput larval development assay, synchronized l1 larval stage animal, selfing caenorhabditis animal, caenorhabditis animal, molecular devices imagexpress nano micropscope, development of animal, selfing caenorhabditis, htlda, quantitative measurement
Funders Acknowledgements:
National Institutes of Health
Abstract
The high-throughput larval development assay (HTLDA) is used to prepare selfing Caenorhabditis strains and quantitatively measure the development of animals using the Molecular Devices ImageXpress Nano micropscope. Synchronized L1 larval stage animals are grown for 48 hours and then measured for animal length as a proxy for development. The HTLDA can be scaled to measure the development of up to 96 unique strains at a time, which provides an efficient method to obtain quantitative measurements of selfing Caenorhabditis animals.
Materials
Some equipment and protocols are described in more detail on the Andersen lab webpage (www.andersenlab.com/protocols)

Equipment:
  • Eppendorf 5810R
  • Plastic shoe box (IRIS USA CNL clear Latching Box, 6 Qt, 18 Count; or similar box)
  • iButtons (Embedded Data Systems, Cat # DS1925L-F5)

Consumables:
  • 12-well V-Bottom Reservoir (USA Scientific, Cat # 3824*3412)
  • 96-well culture plates (USA Scientific 96 well CELLSTAR Clear TC PLT*FLAT, Cat # 5665-5180)
  • Gas permeable sealing film (Fisher, Cat # 14-222-043)
  • Kanamycin (US Biological, Cat # K0010)
  • Sodium azide
  • Bleach solution

Bleach solution (per 200 mL):

Note: All components of the bleach solution should be made four days before use.

1) Make 10 mL of 10 M sodium hydroxide (NaOH):
1a) Add 4 g NaOH pellets to an autoclaved glass media storage bottle.
1b) Measure 10 mL of ddH2O in a graduated cylinder and add the 10 mL of ddH2O to the 4 g NaOH pellets.
2) In a graduated cylinder, measure out 40 mL NaOCl (Fisher, Cat # SS290-1). Do not use store-bought bleach.
3) Add ~ 100 mL ddH2O to the graduated cylinder with NaOCl.
4) Pour the 140 mL of NaOCl + ddH2O solution to the glass media storage bottle.
5) Add ~ 50 mL ddH2O to the graduated cylinder to rinse out the remaining NaOCl and add to the glass media storage bottle. 6) Mix well and store in the dark at 4ºC until needed.
K medium (per 500 mL):

1) Combine the following into a graduated cylinder:
  • 51 mM NaCl (5.1 mL of 5 M NaCl)
  • 32 mM KCl (16 mL of 1 M KCl)
  • 3 mM CaCl2 (1.5 mL of 1 M CaCl2)
  • 3 mM MgSO4 (1.5 mL of 1 M MgSO4)

2) Mix the four salts with increasing water, then fill to 500 mL of ddH2O.
3) Filter sterilize with 0.2 µm Thermo Scientific Filter Unit (Cat # 566-0020).
4) Add 1.25 μg/mL unfiltered cholesterol (125 μL of 5 mg/mL cholesterol) to the K medium after filtering. 5) Mix, label, and store for up to two weeks.

Note: Be sure to check the K medium before use for any floating particulate, which is a sign of contamination. If K medium is contaminated, make new K medium.

M9 buffer (per 1 L):

  • KH2PO4 3 g
  • Na2HPO4 6 g
  • NaCl 5 g
  • ddH2O up to 1 L

1) First measure all reagents into 750 mL distilled water in a 2 L beaker and fully dissolve. Then bring volume to total of 1 L in a graduated cylinder.
2) Filter sterilize using the vacuum filtration systems into 500 mL bottles (2 bottles total) (Thermo Scientific 0.45 µm, Rapid Flow PES, Fisher Cat # 09-740-63B).
3) Autoclave on liquid cycle, 30 minutes (Make sure to loosen the cap only a little, to maintain final volume).
4) Add 0.5 mL of filter-sterilized MgSO4 to 500 mL of M9 buffer.

Sodium azide solution:

Add 6.5 g sodium azide to 2 L 1X M9 buffer.
Section 1: Growing Strains
6d

Note
General notes on amplifying strains:
  • Ensure that your source NGMA (Nematode Growth Medium Agarose) starved plate is no more than one month old before chunking. If your NGMA starved plate is older than one month, generate a newly starved source plate to use in the assay.
  • All source NGMA plates should be stored at15 °C .
  • For picking each generation, use 6 cm plates from the same batch of NGMA and OP50 to account for variation in NGMA and E. coli effects on the assay.
  • Ensure 6 cm NGMA plates are put at room temperature the night before picking so that you will not be picking to cold plates that have condensation.
  • Organize 6 cm NGMA plates containing the same strain into the same cardboard storage box when possible, stacking no higher than five plates high. Whenever possible, organize groups of strains into cardboard boxes such that similar strain names (e.g., JU3291 vs. JU2391) are not in the same box to avoid potential strain confusion. Also, mark plates with similar strain names so that researchers will more easily notice the differences between the strain names.
  • It is NECESSARY to monitor your strains throughout the whole HTLDA process. If strains are growing slowly, you may leave the worms at 21.5 °C (or higher) the day and/or the night before you pick them. Animals (Caenorhabditis elegans and Caenorhabditis briggsae) are reared under standard conditions, including generation zero of the HTLDA. Start monitoring at the first generation to make subsequent picking as easy as possible. If animals are growing faster or slower, they can be moved to a lower or a higher temperature, respectively, for a generation or two. Be careful of temperature-sensitive strains or mutants, because they are most affected by shifting temperatures.
  • For Caenorhabditis tropicalis, all steps should be performed at 25 °C .
  • Monitor the temperature in all incubators to ensure that the incubators match the temperature in the HTLDA protocol. iButtons (or similar temperature-tracking devices) should be used for temperature monitoring in all experiments.




Using a sterile, autoclaved spatula, chunk from a starved NGMA "source" plate onto two OP50-spotted 6 cm NGMA plates. Store these 6 cm NGMA "chunk" plates in a 20 °C incubator.

Note
Ensure your starved NGMA source plate is no more than one month old before chunking. Aim to chunk a plate that yields about 50 adult worms, assessed by the number of arrested larvae on the source plate.

After 48:00:00 , pipette 15 µL of bleach solution (see Materials for recipe) on a new 6 cm NGMA plate away from the E. coli lawn. Pick ten to twenty gravid hermaphrodites from the chunk plate into the bleach spot. Swirl gently with the pick to ensure all animals are removed from the pick. Store these 6 cm "spot bleach" plates lid side up in a 20 °C incubator.

2d
After 24:00:00 , pick 20-30 L1s from the spot bleach plate to a new labeled 6 cm NGMA plate for each strain. If the L1s did not survive the bleach, make a note of it so that larger chunks or fresher source plates can be used for future assays. This pick is generation one. Store these 6 cm "generation one (G1) L1" NGMA plates in a 20 °C incubator.

Note
Keep your spot bleach plates along with your G1 L1 plates in case you need to go back to them in Step 5.

1d
After 48:00:00 , pick five late-stage L4 hermaphrodites from the G1 L1 plates (or spot bleach plates, depending on numbers of animals) to the labeled 6 cm NGMA plates for each strain. These animals are distinguished by a distinct white half-moon shaped patch near the midsection of the animal, containing what looks like a classic "Christmas tree" where the vulva is developing. These plates are "G1 L4" plates. Pick at least three 6 cm G1 L4 plates per strain. Grow this generation of animals at 20 °C .

Note
Animals might have grown into young adults by this time, in which case, pick the youngest adult animals you can find. If you find males on the G1 L1 plates, avoid picking young adults from these plates. Check the G1 L1 plates one and two days after picking to ensure that they will be properly staged for the generation two pick.

Note
Depending on how many generation two (G2) L4 plates are needed in Step 6, you might need to modify the number of G1 L4 plates that are picked. Here are two examples of how you can determine how many G1 L4 plates to pick:

Example 1: If you need to pick five NGMA plates per strain for your G2 L4 pick, pick at least three G1 L4 plates. Even though it may be possible to obtain five G2 L4 plates worth of animals from one G1 L4 plate, picking three G1 L4 plates here safeguards against your single plate becoming contaminated or containing males at the next step, ending your assay.

Example 2: If you need to pick 36 plates per strain for your G2 L4 pick, pick at least five G1 L4 plates in this step.

After monitoring growth rates, pick five late-stage L4 hermaphrodites (look for the developing vulva in the crescent) of each strain to each plate for this generation. Typically, this pick is less than 72:00:00 after the G1 L4 pick. This pick is the G2 L4 pick (AKA the “big pick”). Try to pick as fast as possible. You do not want one strain to be four hours older than the last strain. Do not pick males and avoid picking L4 hermaphrodites from plates with males if other plates have enough hermaphrodites. Grow the G2 L4 pick at 21.5 °C . Check the 6 cm G2 L4 plates for L2 and L3 larvae after two days and L4 and young adults after three days. If animals are L1 or early L2 larvae after two days, then incubate plates at a warmer temperature.

Note
If you predict that the G2 L4 pick will take more than four hours, split up the picking between multiple people to speed up the picking process.

Based on calculations from previous experiments, each G2 L4 6 cm plate provides approximately 1,000 embryos for the average C. elegans strain (using bleach solution). Based on your experimental requirements, calculate the number of plates that you need to pick.

For example:

An assay with eight strains and twelve conditions for each strain with three replicate bleaches and four technical replicates per condition would require:

1) 12 conditions x 4 technical replicates x 3 bleaches/assays x 30 embryos = 4,320 embryos
2) 4,320 embryos / 1,000 embryos / 6 cm plate = Five 6 cm G2 L4 plates (rounded up)

Therefore, for this example experiment, you should pick five 6 cm G2 L4 plates per strain to meet the experimental requirements.

In subsequent experiments, if you find that you are producing a vast excess or dearth of embryos compared to your experimental requirements for your strain(s) of interest, you should adjust the number of picked plates accordingly. The above calculations provide an approximate starting point.

3d
In the afternoon of the G2 L4 pick (i.e., four days before bleaching or ~84-90 hours), make fresh bleach solution for your assay (see recipe in Materials section). Store bleach solution in the dark at 4 °C until needed.

In the afternoon of the G2 L4 pick, make fresh K medium for your assay (see recipe in Materials section).

At any point in the assay, if you notice your K medium has gone bad (any solute crashing out and appearing as white flakes or granules floating), make a new batch.
Section 2: Preparation of synchronized animals
45m
Four days after picking the G2 L4 plates, check the strains to ensure your worms are staged properly for the bleach. Plates are staged properly for the bleach when the population on the plate comprises mostly gravid adults, and adults have begun to lay embryos throughout the plate and have begun migrating away from what remains of the bacterial lawn.
00:45:00 before you start washing your 6 cm plates (step 11), remove the bleach solution from 4 °C and allow it to come to room temperature (keep it in the dark). Room-temperature bleach more effectively dissolves adult worms.

Note
Because of the variation between bleaches, it is highly recommended to perform multiple (three) bleach synchronizations per day. Make sure to pick enough 6 cm plates in the G2 L4 pick and make sure your synchronizations are performed independently (not at the same time). If possible, bleach all strains for an assay together. If it is not possible, bleach the same set of strains together for each bleach-assay. Independent bleach synchronizations comprise separate preparations of bleach solution and performed three separate times.

Follow instructions for conical bleaching in Steps 11 - 22.
45m
Wash worms off plates into a labeled 15 mL conical tube. Dispense M9 buffer onto one 6 cm plate of Strain A, transfer (by decanting) the liquid to a second 6 cm plate of Strain A, transfer the liquid to a third 6 cm plate of Strain A, and finally transfer the liquid to a fourth 6 cm plate of Strain A. Wash off a maximum of four plates from one strain at a time.

Wash the same 6 cm plates of Strain A a second time, pouring M9 buffer onto the last 6 cm plate and transferring M9 buffer to each plate in the opposite direction as was done the first time.

Pour all M9 buffer plus worms into one labelled 15 mL conical tube per strain.

Note
It is easy to pour from the final 6 cm plate into the 15 mL conical with a little bit of practice. We typically pour ~ 2 mL of M9 buffer onto one plate and transfer the same solution to four other 6 cm plates before pouring it into the conical. Repeat for all strains.


Spin down worms in 15 mL conical tubes at 254 g (Eppendorf 5810R, 1100 rpm) for 1 minute using a table-top clinical centrifuge.

Aspirate off the M9 buffer, being careful to not aspirate any of your worms.

Once all strains are pelleted in a 15 mL conical tube and all M9 buffer is aspirated, proceed to the next step.

Add 7 mL of bleach solution to each 15 mL conical for pellet sizes of 100 µL and greater. If pellet sizes are smaller, adjust volume of bleach solution accordingly. We use 4 mL of bleach solution for all smaller pellets.

Note
The freshness of the bleach solution matters. Ineffective bleach solution (e.g., kept at 4 °C longer than one month or kept at room temperature for longer than eight hours) will require more time to bleach worms and will not dissolve the worms uniformly.


Shake the tubes manually and vigorously. If multiple people are bleaching, try to ensure that each individual is shaking in approximately the same way to ensure greater consistency between strains bleached by multiple individuals.

After three minutes of shaking, check every 30 seconds to see if the adult worms are dissolved. Be very careful not to over-bleach the worms. Once you see that the carcasses are nearly dissolved, move to the next step. If only embryos are left, then you have gone too long. If you find a strain that is lagging, shake this strain more vigorously to catch it up to the other strains.

Note
We have found it useful to shake to the beat of a song. Use a playlist with songs that have the same beats per minute (bpm). We use 130 bpm. Shake by holding the tubes by the cap to avoid the transfer of warmth from the hands to the solution. You can bleach multiple strains at the same time, but depending on experience, you may want to start with only doing 4-5 strains at once. The most you should bleach at a time is 10 strains.

Once you find that only a few adults remain, move quickly to spin down the embryos at 254 g (Eppendorf 5810R, 1100 rpm) for 30 seconds. It is best to have the reagents next to the waste receptacle and prepared for the next few steps.

Note
Some table-top centrifuges do not have an option of 30 seconds, so you must stop the centrifuge manually after 30 seconds. If you feel you have under-bleached your worms, you may let them spin down for a full minute.

Immediately decant off the bleach into the waste receptacle carefully and quickly without disturbing the worm pellet. It is critical to move as fast as possible from Step 16 to Step 21.
Pour 10 mL of M9 buffer from a 50 mL conical tube to each 15 mL conical tube of worms.

Invert three times as you walk back to the clinical centrifuge.
Spin down the embryos at 254 g (Eppendorf 5810R, 1100 rpm) for 30 seconds.
Decant off the M9 buffer into the sink or waste container carefully and quickly without disturbing the worm embryo pellet.
Repeat steps 16-20 twice with M9 buffer.
Pour 10 mL of K medium from a 50 mL conical tube to each 15 mL conical tube of worms.
Invert three times as you walk back to the clinical centrifuge.
Spin down the embryos at 254 g (Eppendorf 5810R, 1100 rpm) for 30 seconds.
Decant off the K medium into the sink or waste container carefully and quickly without disturbing the worm embryo pellet.
After the final wash, resuspend the embryos in 3 mL of K medium. Adjust the volume of K medium to the number of plates that you bleached. For 15 6 cm plates per bleach, we find 3 mL is a good volume. For fewer 6 cm plates, 2 mL might be more appropriate.

Determine the titer of the embryos by counting the number of embryos in five replicates of 3 µL onto a lid of a 96-well plate wiped with Tween 20.

Pulse vortex, by starting and stopping the vortex every few seconds, each 15 mL conical tube of embryos for at least ten seconds to ensure embryos are homogeneously dispersed in the K medium.

Note
If embryos are too numerous (more than 50 per droplet) to count, you should dilute the embryo suspension further to ensure accurate counts.

If embryos are too dilute (fewer than 5 per droplet), spin down the embryos for 1 minute at 254g (Eppendorf 5810R, 1100 rpm) and carefully pipette off some of the K medium and repeat the embryo counts.


Adjust the resuspension liquid (K medium) to obtain the desired concentration (0.6 embryo per μL).
Pulse vortex each 15 mL conical tube of 0.6 per µL embryos for at least ten seconds before pouring into a sterile reservoir.
Using a multi-channel pipette, pipet immediately and ensure proper mixing before pipetting into assay plates. To mix, take 50 µL from the bottom of the reservoir and dispense it higher up on the side of the reservoir, repeat on the other side for a total of ten times (five per side).

Pipet 50 µL of embryo solution into each well of the tissue-culture treated 96-well flat-bottom plate (USA Scientific 96 well Cellstar Clear TC PLT*FLAT Cat #: 5665-5180) to dispense approximately 30 embryos per well.

Note
Example: With 12-channel reservoirs, you can prepare a row of each 96-well assay plate in each reservoir.


Seal all 96-well plates with a gas-permeable sealing film (Fisher, Cat # 14-222-043).

Try to avoid wrinkles in the film and use a fingernail to seal edges of the plate and wells.
Place all 96-well plates in a humidity chamber.

To make a humidity chamber, place paper towels saturated with ddH2O in a clean (bleached with 10% solution of household bleach, 70% ethanol rinsed, and ddH2O rinsed) plastic box (IRIS USA CNL clear Latching Box, 6 Qt, 18 Count; or similar box).
After filling the humidity chamber with plates, put on the lid and seal the chamber using parafilm.

Note
These chambers hold 18 plates, so scale the number of boxes according to your experiment.

Shake overnight at 170 rpm at 20 °C (Excella E24R shaker or the INFORS HT Multitron shaker) in the humidity chamber.

Section 4: Feeding strains
3d 18h
To keep your 96-well plates clean and debris-free, conduct the following steps:

A) Approximately 30 minutes prior to feeding strains, place an air filter on the bench where you plan to feed strains. Keep this air filter on during the feeding process.

B) Put on fresh gloves, a face mask, and a lab coat. For researchers with long hair, tie your hair back.

C) Clean your bench and all used supplies with a 10% solution of household bleach followed by 70% ethanol.
The morning after bleaching, check one or two 96-well plates that were aliquoted the previous day. Inspect the plates for successful hatching of L1 larvae under a stereoscope.

Nicely swimming L1 larvae suggest successful bleach synchronization, so continue with protocol. Also, check for leftover chunks/contamination from the bleach and crystals. You want the wells to be as clean and contamination-free as possible. Crystals often arise if M9 buffer is not washed out by K medium. If you observe any issues noted above, document the well(s) for the data processing steps.
If you are testing the effects of drugs or toxicants on worms, prepare drug dilutions and add to OD30 HB101 food with kanamycin.
If you are testing the effects of drugs or toxicants on worms, ensure you also prepare the appropriate vehicle control conditions for those drugs or toxicants (whatever diluent is used for your drug, toxicant, etc.). Typically, we use either ddH2O or DMSO.
Make food from previously prepared, frozen E. coli HB101 OD100. See other Andersen lab protocols for bacterial preparations.

If your assay contains many 96-well plates, start making food early in the morning.


Note
We recommend using the Iterative Batch Averaging Method (IBAT) to make your OD30 HB101 (Step 35.1) to feed the animals in the HTLDA. IBAT produces a consistent food source that can be used across HTLDAs.

For example, if 10 mL of HB101 OD100 food is needed to feed animals in 96-well plates, take one mL from ten unique HB101 food preps, allowing you to average food prep effects across ten batches.

IBAT reference: https://pubmed.ncbi.nlm.nih.gov/34156835/

Dilute freshly thawed OD100 HB101 to a concentration of OD30 with K medium. This dilution should be made in an autoclaved Pyrex media storage bottle.
Add kanamycin (US Biological, Cat # K0010) to the OD30 food mix. The final kanamycin concentration in the well should be 50 μM (final concentration in the food dilution, prior to plating, will be 150 μM).

18.75 µL of 80 mM stock per 10 mL of solution

Start with one or two 96-well plates. Make sure to resuspend the HB101 food by swirling (if a large volume of OD30 HB101 is made in an autoclaved Pyrex media storage bottle) or vortexing (if using smaller batches of OD30 HB101 made up in a conical tube or otherwise vortexable container) before you add it to the 96-well plates.
The following steps will describe how to feed your 96-well plates.

Your goal is to feed one 96-well plate on average every 3-4 minutes to match the time for imaging on the Molecular Devices ImageXpress Nano. Use a timer to make sure that you do not forget. Ensure that the HB101 food is mixed into each well. Every well should look cloudy.

Note
It is easiest to see which wells have been fed when you look at them from above and on a black background.

Mix the volume of HB101 and drug (drug is optional) required for three replicate 96-well plates. Pour this mixture into a single-channel reservoir (USA Scientific, Cat # 1306-1010).

Using a micropipette, add 25 µL of food (and drug) to every well containing worms for each of the replicate plates.

Seal with a gas permeable sealing film (Axygen, Fisher Cat # 14-222-043), label the plate top with the plate number and any condition information, and record the time that you added the food solution.
Put the 96-well plates back in the humidity chamber once they are fed. When done for the day, make sure your paper towels are still damp and seal the chamber with parafilm. Return the humidity chamber to the incubator.
Shake for 48:00:00 at 180 rpm at 20 °C (Excella E24R shaker or the INFORS HT Multitron shaker) for C. briggsae and C. elegans. Shake for 42:00:00 at 180 rpm at 25 °C for C. tropicalis.

3d 18h
Section 5: Scoring animals
3d 18h 10m
EXACTLY 48:00:00 (C. briggsae or C. elegans) or 42:00:00 (C. tropicalis) after a 96-well plate was fed and drugged, add 167 µL of 50 mM sodium azide solution (see Materials for recipe) to each well twice 00:10:00 before scoring.

3d 18h 10m
Use a single-channel reservoir to hold the sodium azide solution and dispense it using either an 8- or 12-channel pipette. For each step, dispense at opposing angles to each well.

Note
Opposing angles help alleviate heterogeneous swirling of well contents and uneven illumination of images. Also, incubating in sodium azide solution for too long (> 15-20 minutes) might cause uneven illumination as the bacteria can clump.

Follow your lab's protocol to image each 96-well plate on the ImageXpress.