Chromatin Accessibility Profiling in Whole Caenorhabditis elegans L4 Larvae

Summary

Here, we present a quick and simple ATAC-seq protocol to be implemented in L4-stage Caenorhabditis elegans whole-worm using only 30 µL of worm pellet.

Abstract

Chromatin accessibility plays essential roles in transcription, DNA repair, and chromosome segregation. Hyper-accessible regions usually correlate with active promoters and enhancers, facilitating transcription factor binding and regulatory activity. The assay for transposase-accessible chromatin using sequencing (ATAC-seq) enables genome-wide profiling of chromatin accessibility with very few cells. However, its implementation in Caenorhabditis elegans is limited by the nematode’s rich collagen cuticle that complicates cell dissociation. Here, we present an optimized protocol for performing ATAC-seq in whole worms at the L4 stage. The procedure begins with synchronized cultures and involves cuticle disruption, enzymatic dissociation, and cell-suspension preparation. Permeabilized nuclei are then subjected to Tn5 transposition, followed by polymerase chain reaction (PCR) amplification and purification of next-generation sequencing (NGS)–ready libraries. This protocol requires 30 µL of worm pellet, can be completed in one day, and generates 5,000–9,000 accessibility peaks in the Bristol N2 reference strain. This streamlined workflow can be adapted to other developmental stages or FACS-purified cell populations. By reducing technical barriers to ATAC-seq in C. elegans, this method expands opportunities to study genome-wide chromatin accessibility in response to genetic and environmental perturbations in a whole-organism context.

Introduction

Eukaryotic genomes are densely packed inside the cell nucleus. To cope with spatial constraints, DNA is tightly wrapped around nucleosomes, multiprotein complexes composed of eight histones that are highly conserved across different phyla¹. Chromatin accessibility patterns vary depending on the cellular state in a tissue, and gene-specific patterns^2,3,4. Nucleosome accessibility strongly correlates with active transcription, particularly at promoters and distal regulatory regions such as enhancers, which are accessible to transcription factors⁵. Open chromatin regions also play important roles in DNA repair, replication, and chromosome segregation³. In contrast, reduced accessibility usually correlates with silenced heterochromatin and repetitive sequences^6,7.

Mapping chromatin accessibility has been a powerful approach for understanding gene regulation. Different techniques have been employed to interrogate chromatin accessibility. Among the earliest methods, micrococcal nuclease (MNase) and DNase I digestion were used to map nucleosome positioning and accessibility at specific loci^8,9. These techniques were later implemented for genome-wide studies when coupled with next-generation sequencing (NGS)^10,11. However, these techniques require careful calibration of enzyme concentration and reaction time, along with a substantial number of cells, for successful implementation.

In 2013, the Greenleaf laboratory established the assay for transposase-accessible chromatin using sequencing (ATAC-seq) to seamlessly map chromatin accessibility at a genome-wide level using a low number of cells¹². ATAC-seq takes advantage of the commercially available Tn5 transposase to preferentially insert short DNA sequences (i.e, barcodes) into open chromatin regions, typically devoid of nucleosomes, and correlates with active promoters and enhancers. The protocol requires cell permeabilization and nuclei isolation, followed by limited digestion with Tn5 for tagmentation of hyper-accessible chromatin. Later, complementary indexes are incorporated during PCR, facilitating sample multiplexing before NGS¹³. ATAC-seq has proven to be a robust protocol that also enables in silico transcription factor footprint and has been implemented for single-cell resolution^14,15,16,17. In addition to mice and human samples, ATAC-seq has been implemented in various systems, including zebrafish and invertebrates such as Drosophila and the nematode Caenorhabditis elegans^18,19,20.

For decades, C. elegans has been a powerful model system in fields such as development, cell biology, and neuroscience, leading to seminal discoveries including the identification of micro RNAs and the RNA interference (RNAi)^21,22. It has a compact 100 MB genome with ~20,000 protein-coding genes²³. Gene regulation occurs through promoter activity as well as distal enhancers, which have been mapped via ATAC-seq across different tissues, developmental stages, and aging time points^24,18. Chromatin accessibility profiling has thus become a valuable tool for understanding gene regulation in C. elegans. Consequently, ATAC-seq holds particular promise for studying epigenetic landscapes across different environmental conditions or wild isolates to uncover diverse transcriptional responses. Additionally, it could provide insights into the chromatin architecture of other pathogenic and non-pathogenic nematodes.

One of the primary methodological challenges in C. elegans is the presence of a collagen-rich cuticle that must be dissociated to obtain a homogenous cell suspension from whole worms²⁵. To address this challenge, we compiled and optimized different protocols to disintegrate the nematodes and then perform ATAC-seq in C. elegans^18,26. Using the L4 larval stage is preferred as animals at this developmental point possess a transcriptionally active germline, which translates into open chromatin, providing a consistent and informative source of chromatin accessibility in somatic cells. Here, we present a rapid and efficient protocol for worm dissociation and cell suspension generation, enabling ATAC-seq in C. elegans. Our method is optimized for L4-stage worms and requires as little as 30 µL of worm pellet, with the potential to be adapted for other developmental stages or for smaller samples, such as FACS-purified cells. This protocol can be performed in different C. elegans strains and conditions, enabling the identification of 5,000–9,000 chromatin accessibility peaks in the Bristol-N2 reference strain, laying the foundation for subsequent chromatin accessibility analyses.

Protocol

It is recommended to wear gloves at all times.

NOTE: Day 1: Preparing the synchronized worm population. All worms were grown on NGM plates seeded with E. coli OP50. Things needed before starting: (per sample or strain), One 90 mm or two 60 mm adult worm plate(s), One 90 mm or two 60 mm plates with food, M9 sterile buffer²⁷ (3 g KH₂PO₄, 6 g Na₂HPO₄, 5 g NaCl, 1 ml 1 M MgSO₄, H₂O to 1 L. Sterilize by autoclaving)

1. Collection of worms

1. Take a 90 mm (or two 30 mm) plate with the worm’s adult population.
2. Recover the worms with M9 buffer and transfer them into a 15 mL centrifuge tube.
3. Repeat the previous step to recover all the worms from the plate(s).
4. Let the worms settle in the 15 mL tube for a minute and carefully remove the M9 buffer.
5. Be sure to have ~1 mL of worms.

2. Bleaching

1. Add 5 mL of M9 buffer + 650 µL of NaOH 5M + 1,300 µL Household bleach (5% solution of NaClO).
2. Vortex at max. velocity for 1–2 min (not more than 2.5 min).
3. Centrifugate at 1,000 x g for 1 min at RT (room temperature).
4. Remove the supernatant and add 9 mL of M9 buffer.
5. Repeat sections 2.3 and 2.4 five more times.
6. Resuspend the embryos in 7 mL of M9 buffer.
   NOTE: Bleaching protocol may vary in efficiency depending on temperature, type of commercial bleach²⁸.

3. M9 (liquid) overnight hatching of the eggs

1. Be sure that the collection tube with the bleached eggs is properly closed.
2. Put the tube in slow movement at 20 °C (e.g., in a shaker inside a room at 20 °C).
3. Worms will hatch overnight and arrest in L1.
   NOTE: Don’t leave the tube for more than 12 h.

4. Day 2: growth of worms: recovering L1 worms.

1. Take the tube from the shaker and corroborate under the microscope that there are L1 worms.
2. Let the tube settle vertically for about 1 h at RT.
3. Remove the supernatant and transfer ~300 L1 worms to one 90 mm or two 30 mm plates with food.
4. Place them in a 20 °C incubator and let them grow for 26 h. to reach the L4 stage.

5. Day 3: Preparation of stock solutions: The next solutions must be prepared to make fresh solutions on day 4

NOTE: This can be done on day 2 or 3.

1. 1 mL of 10% SDS.
2. 50 mL of 1M HEPES pH 8 (autoclave).
3. 250 mL of 20 % Sucrose (autoclave).
4. 100 mL of 2M NaCl: prepare (autoclave).
5. 50 mL of 2M KCl (autoclave).
6. 10 mL of 1M CaCl₂ (autoclave).
7. 10 mL of 1M MgCl₂ (autoclave).
8. 100 mL of 1M Tris, pH 7.4.
9. 5 mL of 5M NaCl (autoclave).
10. 500 µL of 10% Tween-20.
11. 500 µL of 10% Nonidet P40/IGEPAL.
12. 100 µL of 5% Digitonin (needs to be heated to 65 ˚C to dissolve).
13. 500 µL of 10% BSA (filter with 0.22 µm) stored at 4 ˚C.
    Day 4: L4 stage worms for ATAC
    NOTE: Before starting, be sure to have an incubator at 25 ˚C, a 4 ˚C centrifuge ready, heat up Digitonin to 65 ˚C (for fresh buffer preparation), have sterile M9 buffer, sterile H2O, and 20 µm filters.

6. Solution preparation

1. Prepare fresh solutions (see Table 1). Always handle L-15 10% FBS and egg buffer in sterile conditions and keep them cold.

Table 1: Stocks for fresh solution preparation. Recipes for stock preparation for nematode dissociation and cell suspension protocol (sections 6 and 7). Please click here to download this Table.

7. Nematode dissociation and cell suspension protocol

NOTE: One 90 mm or two 30 mm plates with an L4 synchronized worm population will be needed.

1. After 26 h in the 20 °C incubator, corroborate under the microscope that the majority of the worms are in the L4 stage.
2. Recover all the worms from the plates with M9 buffer and put them into a 1.5 mL centrifuge tube.
3. Wash 5 times with 1 mL of sterile H₂O by letting the worms settle by gravity.
4. Transfer the tube with the worms to a 25 °C incubator for 30 min to get rid of intestinal food remains.
5. Take 50 µL of the worm pellet and place it into a new tube.
6. Add 200 µL of SDS-DTT fresh buffer and mix in a tube rotator for exactly 5 min (not longer).
7. Add 1 mL of cold egg buffer and give a spin on a picofuge to remove supernatant. Repeat 6 times.
8. Add 80 µL of fresh prepared pronase (15 mg/mL).
9. With a 200 µL pipette (with filtered tips), mix 70 times the worms-pronase mix and incubate for 15 min at RT.
10. Add 800 µL of L-15 10% FBS media and centrifuge 5 min at 500 x g at 4 °C.
11. Add 1 mL of L-15 10% FBS media and repeat centrifugation.
12. Repeat the previous step twice more.
13. Remove supernatant and re-suspend in 900 µL of L-15 10% FBS media.
14. Filter with a 20 µm filter.

8. Nuclei isolation and Chromatin tagmentation

NOTE: Use pipette filtered tips. Before starting: Take out Tn5 Buffer, let it thaw, set a thermoblock to 37 °C, take out PCR product purification kit (Table of materials), put EB Buffer at 37 °C, Centrifugation of the columns should be performed at RT.

1. Centrifuge the filtered cell suspension for 5 min at 500 x g at 4 °C.
2. Remove supernatant, leaving ~50 µL.
3. Add 45 µL of Lysis Buffer (freshly prepared) and carefully re-suspend.
4. Carefully add 50 µL of Wash Buffer (do not mix).
5. Centrifuge for 5 min al 500 x g at 4 ˚C. Remove supernatant. Keep it in the ice.
6. While spinning, prepare for each reaction (Rx) the following Tn5 mix per sample: 25 µL Tn5 Buffer²⁹, 24.8 µL Nuclease-free H₂O, 0.2 µL Tn5 enzyme. Prepare a master mix for multiple samples.
7. After centrifugation, remove the supernatant. Keep the nuclei on ice. Add 50 µL of Tn5 mix to the nuclei pellet and slowly pipette seven times. Incubate at 37 °C for 30 min on a thermoblock. Make sure to give little taps to each tube every 10 min.
   NOTE: Remove the PCR purification kit and place the EB Buffer in a 37 °C incubator. The next 5 steps will use this Kit’s buffers.
8. After Tn5 incubation, add 300 µL of PB buffer (Yellow) from the kit and mix in a vortex at maximum speed for 25 s.
9. Take all the volume from the tube and transfer it into the spin column (purple, stored at 4 °C) and centrifuge 1 min at maximum speed at RT.
   NOTE: The column will hold the DNA, so the supernatant can be trustingly discarded.
10. Add 750 µL of PE Buffer to the spin column and centrifuge 1 min at maximum speed at RT.
11. Discard the supernatant with the collection tube. Place the column in a new 1.5 mL tube and repeat centrifugation; discard the 1.5 mL tube.
12. Place the column in a new, labeled 1.5 mL tube and add 10 µL of EB Buffer at 37 °C to the center of the column. Be sure not to touch the column. Leave it at RT for 1 min.
13. Centrifugate for 1 min at maximum speed at RT; never at 4 °C.
14. Repeat the last 2 steps for a total of 20 µL elution in EB buffer.
    NOTE: Optional pause: If stopping, sample(s) can be stored at -20 °C. Here, the tagmented chromatin is in a 20 µL volume.

9. Library amplification

1. For ATAC library amplification, a real-time PCR has to be performed to estimate the proper cycle amplification number (see Table 2). Perform this on a PCR plate.
2. To estimate the cycle numbers for final amplification, the cycle number that corresponds to the first third of the amplification curve has to be taken. It is ideal for the cycle number not to exceed 15. The cycle number must be estimated for each sample.
   NOTE: After qPCR, the tagmented DNA is in a 18 µL volume. Optional pause: If wanting to stop, samples can be stored at -20 ˚C.
3. ATAC Library final amplification:
   1. Each sample must have a different Rv primer (take notes).
   2. Each sample must be amplified for the cycle number (N) estimated for each one.
   3. Perform this PCR in 200 µL tubes (see Table 3).
   4. Here, there are 8 µL of tagmented DNA and 50 µL of amplified ATAC libraries
   5. Optional pause: If wanting to stop, sample(s) can be stored at -20 ˚C.

Table 2: Instructions for first library amplification. Recipe and PCR program for first library amplification (section 9). Please click here to download this Table.

Table 3: Instructions for ATAC Library final amplification. Recipe and PCR program for final library amplification (section 9). Please click here to download this Table.

10. Library purification

NOTE: Before starting, take the size selection beads (AMPure, SPRIselect, or homemade calibrated beads) out of the fridge to let them adjust to room temperature. Mix the beads vigorously using the vortex every time before using.

1. Prepare 1 mL of 80% Ethanol (molecular grade).
2. Add 25 µL (0.5 vol) of beads to the sample tube (where there are 50 µL) and mix with the pipette.
3. Incubate at RT for 10 min.
4. Place the tubes in the magnetic rack for 5 min.
5. Transfer the supernatant (70 µL) to a new clean 200 µL tube.
   NOTE: In this step, the <200 bp DNA fragments are discarded.
6. Add 65 µL (1.3 vol) of beads and mix 10 times.
7. Incubate at RT for 10 min.
8. Place the tubes in the magnetic rack for 5 min.
9. Discard the supernatant (without taking the tube from the magnetic rack).
   NOTE: At this point, the beads are holding the >200 bp and <1000 bp DNA fragments.
10. With the tube still on the magnetic rack, add 200 µL of fresh prepared 80% ethanol directly over the beads and pipet up and down several times to wash them.
11. Remove ethanol and repeat washing with 200 µL of ethanol.
12. Discard the ethanol, then let the beads dry with the lid open for ~2–5 min (don’t let them dry too much).
13. Take the sample(s) out of the magnetic rack. Re-suspend in 50 µL of sterile H₂O. Make sure to wash all beads from the tube's walls.
14. Finally, incubate the tube(s) in the magnetic rack for 5 min.
15. Transfer the supernatant to a new clean tube. Here, the amplified and purified ATAC Libraries are stored.
    NOTE: Libraries can be stored at -20 ˚C for a short time (up to four weeks) or for longer periods at -80 ˚C.

11. Fragment size distribution

1. Once the final amplification is done, it is advisable to run a part of the sample through a capillary electrophoresis to be able to see the fragment size distribution pattern.

Results

Quality control after tagmentation ensures accurate library preparation
During the protocol, it is important to pay special attention to qPCR performed after chromatin tagmentation (section 9), as it is a vital step for assessing DNA quality and the experiment’s performance. Figure 1A shows two amplification curves; the left one (under 15 cycles) represents good quality material that shows that everything was successful until this point, so it is ok to continue. The curve on the right (Figure 1A) shows poor-quality material, so this sample should not be continued with. The next point to pay attention to is that, after amplification and purification of the libraries, it is advisable to run capillary electrophoresis to assess the fragment length distribution. To validate correctly the transposed DNA, fragments corresponding to a nucleosomal pattern should be shown: ~150 bp (mono-nucleosome), ~300 bp (bi-nucleosome), and so on (Figures 1B–D, samples 1 and 2). If the lane shows a smear of fragments, the DNA was overtagmented, and these samples will have to be repeated (Figure 1B and 1E, sample 3).

Sequencing metrics confirm high-quality and scalable library performance
After sequencing, library quality and data yield are evaluated using FastQC. For whole-worm L4 samples, high-quality datasets typically consist of more than 20 million paired-end reads per sample, with duplicate read fractions below 20–30% (Figure 2A). Libraries generated using this protocol are compatible with standard short-read sequencing platforms, such as those from Illumina, and perform robustly with paired-end read lengths of 75 bp. At this sequencing depth, fragment length histograms consistently recapitulate the expected nucleosomal periodicity (Figure 2B), confirming successful chromatin accessibility profiling. Importantly, incorporation of indexed adapters during library amplification enables efficient multiplexing of multiple samples within a single sequencing run, without loss of data quality, making the protocol scalable and cost-effective for comparative studies. Also, Figure 2C quantifies the number of shared peaks between samples 1 and 2, Figure 2D details the annotations of these shared peaks, and Figure 2E visualizes the corresponding read pileup and peak profiles supporting this overlap.

Figure 1: Outcomes for positive and negative results after conducting the ATAC-seq method. (A) Real-time PCR amplification curve. For the first curve (left), the cycle estimation for amplification is under 15, which corresponds to good-quality DNA material. For the second curve (right), the cycle estimation is over 20, which corresponds to a suboptimal quality. (B) Capillary electrophoresis. Samples 1 and 2: the fragment length distribution corresponding to the nucleosomal pattern: ~150 bp (mono-nucleosome), ~300 bp (bi-nucleosome), and so on. Sample 3 shows overtagmentation, indicating that the method did not work. (C and D) Fragment distribution between 150 and 1000 bp corresponding to samples 1 and 2. (E) Fragment distribution corresponding to sample 3, which is the smear of fragments of the failed experiment. Please click here to view a larger version of this figure.

Figure 2: Quality control bioinformatic metrics for two different samples done with this ATAC-seq method. (A) Quality control metrics for samples 1 and 2. For good quality: > 20 M seqs (million sequences); < 20% Dupl (duplicated reads); 35–40% GC. (B) Shows the fragment length histogram that corresponds to the nucleosomal pattern of samples 1 and 2: ~150 bp (mono-nucleosome), ~300 bp (bi-nucleosome), and so on. (C) Number of shared peaks for samples 1 and 2. (D) Shared peak annotations from samples 1 and 2. (E) Pileup and peaks of reads from samples 1 and 2. Please click here to view a larger version of this figure.

Discussion

Here we describe an optimized protocol for profiling genome-wide chromatin accessibility by ATAC-seq in whole L4-stage Caenorhabditis elegans worms. Chromatin accessibility directly influences gene expression, DNA repair, replication, and recombination, and while ATAC-seq has become a standard approach for cell and tissue samples, its application to intact organisms poses unique technical challenges, particularly in nematodes with an impermeable, collagen-rich cuticle. At the same time, assessing whole-organism assays provides a unique opportunity to capture global epigenetic responses. Our method addresses these challenges by refining worm dissociation, permeabilization, nuclei preparation, chromatin tagmentation, and library preparation from small numbers of synchronized L4-stage worms. This protocol can be fully performed in one or two days and allows parallel processing of several samples.

Several steps are critical for the success of this protocol. (1) Freshly prepared buffers and regents are important to reduce sample variability. (2) The protocol is optimized for L4-stage worms from a synchronized population; we expect that the protocol can be adapted to other developmental stages, but worm dissociation should be carefully optimized to prevent cell lysis. (3) During dissociation, the SDS-DTT incubation should not exceed 5 min, and the pronase digestion should be no longer than 15 min; inspection under a microscope is recommended to prevent over-digestion and cell death. (4) In library amplification, it is not advisable to continue with samples that require more than 15 cycles during the first qPCR, as they typically yield low-complexity libraries. (5) The final fragment distribution should ideally display a nucleosomal pattern; although in our experience, libraries with overrepresentation of internucleosomal fragments above 150 bp or showing a flat, wide profile can still yield usable results, but caution is advised. (6) Because the C. elegans genome is 1/30 the size of the human genome, 5–10 million pair-end reads are generally sufficient. (7) ATAC is sensitive to intrinsic and uncontrolled environmental perturbation, generating batch effects; therefore, experimental and control samples should be processed and sequenced together. If sequencing in multiple runs is unavoidable, a representative control should be included in each run for batch correction.

This method is effective with limited worm material (~30 µL worm pellet). If starting with a different amount, reagent volumes during dissociation can be escalated proportionally, but care must be taken not to over-dilute. In our hands, variation in incubation time or temperature during tagmentation was robust across the tested ranges, suggesting flexibility at this step. By contrast, the ratio of nuclei versus Tn5 is critical: excess Tn5 or low cell number promotes over-tagmentation. Overtagmented libraries are typically detected as a single band about 150 bp on electropherograms; in such cases, increasing the input cell number is more effective than reducing the Tn5 exposure/amount.

This protocol is optimized for L4 worms, and although adaptable, each developmental stage will require empirical adjustment, in particular desegregation. Another limitation is the use of whole worms, which yield average chromatin accessibility profiles that mask tissue- or cell-specific signatures. While single-cell ATAC-seq is possible, its implementation in nematodes will depend on the purification of specific cell populations. Given the extensive collection of tissue reporter strains in C. elegans, combining this method with FACS-purified cell populations will enable tissue-restricted chromatin profiling. Future improvements may include the addition of spike-in (e.g., bacterial gDNA or mammalian/Drosophila cells at 1–5%) for normalization of signal across conditions.

Compared with earlier protocols relying on large input material or complex dissociation steps, our method is faster, simpler, and reproducible, making it accessible for laboratories with limited experience in nematode chromatin biology. Importantly, the streamlined workflow allows simultaneous processing of multiple samples, enabling comparative studies of chromatin accessibility under different environmental or genetic conditions. This expands the potential of C. elegans as a model for investigating epigenetic responses to diet, stress, pathogens, drugs, or aging. Whole-animal profiling can provide broad insights into how the organismal epigenome adapts, while the protocol’s adaptability to wild isolates or related nematodes broadens its relevance for evolutionary and parasitology research.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
250 mL, 0.22 µm PES	Ultra Curz	sc-200253
Agar	BD	214010
Agencourt AMPURE XP	Nalgene	A63886
Bovine Serum Albumin	Maplica	200100
Calcium Chloride	Sigma	C5670-100G
CellTrics 20 µm	Sysmex	04-0042-2315
Cholesterol	Sigma	C8503-25G
Digitonin	Sigma	D141-500 mg
Disposable gloves	Microflex	94-243
DL-Dithiothreitol	sigma	D0632-10G	Harmful
Ethyl alcohol, pure	Sigma	1003661421
Fetal Bovine Serum	biowest	S181S-500
HEPES	Sigma	H3375-250 G
Illumina Tagment DNA Enzyme and Buffer (large Kit)	Illumina	20034211
Kapa Syber 50 mL	Sigma	KK4618
L-15 Medium (Leivobitz)	sigma	L4386-10L
Magnesium Chloride anhydrous	Sigma	M8266-100G
Magnesium Sulfate Heptahydrate	Sigma	M1880-500G
Magnesium Sulfate Heptahydrate	Sigma	M1880-500G
MinElute PCR Purification Kit (50 preps)	Qiagen	28004
NEBNext® High-Fidelity 2X PCR Master Mix	NEB	M0541S
Nonidet P40 Substitute	Sigma	74835-1L
Pen Strep	Gibco	15140-122
Peptone	BD	211677
Petri dish 60 mm	Tritech	T3308
Petri dish 90 mm	Interlux	C9015-2C
Pipet tips 10 µL, filter	Cellpro	800108
Pipet tips 20 µL, filter	Cellpro	800708
Pipet tips 200 µL, filter	Cellpro	800608
Potassium Chloride	Sigma	P9541 -500G
Potassium Phosphate Dibasic Powder	J.T. Baker	3252-01
Potassium Phosphate Monobasic, Crystal	J.T. Baker	3246-01 500G
Sodium Chloride	JT Baker	3624-01
Sodium dodecyl sulfate	sigma	L4509-100G
Sodium Hydroxide	Macron	7708-10
Sodium Hypochlorite	Cloralex	544394479
Sodium Phosphate Dibasic 7-Hydrated Crystal	J.T. Baker	3824-01
SPRIselect	Beckman	B23318
Sucrose	Sigma	S0389-500G
Syringe Filter PVDF 0.22 µm	Ultra Curz	Sc-358812
Trizma base	Sigma	T1503 1KG
Tween 20	Sigma	P9416-50ML