How sterile is sterile when working with nucleic acids to prevent contamination?

I am reading up on preparatory work on working with nucleic acids and a lot of the instructions speak on excessive procedures on cleaning environments with high %ethanol and making sure the equipment is nuclease free, and autoclaved.

Are these sterilization steps really necessary when doing research /running gels? Buying all of this equipment seems very excessive, and given that just one nuclease could compromise my results why bother?

Short answer: Yes

Long answer: Depends what you are working with.

DNA: If you are working with DNA, its pretty stable and you can usually get away with a 70% ethanol wash/autoclave (mainly to prevent contamination and obtain consistent results). EDIT: Read Chris's answer also below

RNA: If you are working with RNA well… whatever you did for DNA doesnt apply anymore. You have to take it to the next level. You will need to replace everything, you'll need nuclease free water, tubes, reagents and whatnots. Throw out ethanol and bring in RNAZap, DEPC water or something like that.

An RNase free environment is essential when working with RNA samples. There are two main reasons for RNA degradation during RNA analysis.

First, RNA by its very structure is inherently weaker than DNA. RNA is made up of ribose units, which have a highly reactive hydroxyl group on C2 that takes part in RNA-mediated enzymatic events. This makes RNA more chemically labile than DNA. RNA is also more prone to heat degradation than DNA.

Secondly, enzymes that degrade RNA, ribonucleases (RNases) are so ubiquitous and hardy; removing them often proves to be nearly impossible. For example, autoclaving a solution containing bacteria will destroy the bacterial cells but not the RNases released from the cells. Furthermore, even trace amounts of RNases are able to degrade RNA. Note that RNAses are present everywhere (skin, reagents, normal plastic etc).

Therefore, it is essential to avoid inadvertently introducing RNases into the RNA sample during or after the isolation procedure.

Remember that following microbiological aseptical techniques is usually good practice (and a requirement) when working in a molecular biology lab.

There are excellent guides available online for working with RNA:

I took the above paragraph from here:

Life tech has a few more tips for you:

Here is a good pdf to have in your labbook as a reference sheet:

If this scares you , that's good because when working with RNA there are a million things that can go wrong the first few times you try it. You have to be very careful, and adapting a cavalier attitude such as not bothering is going to come back and haunt you at night…

First of all, sterility is not necessary. It takes much more effort to reach this than just to wipe down everything with ethanol. What you need is a clean and controlled work environment (but this is something you need anyway to get reproducible results) and good and clean equipment. You will need more precautions for RNA work as RNA is more sensitive and RNAses are ubiquituous, but the general rules are the same. Some remarks from me about it:

  • There is no need to wipe down everything in your environment before working. Will you DNA touch the desk or the outside of you pipette? Simply wear gloves, use fresh plasticware, do not touch the plasticware with your bare finger and you will be fine.

  • There is no general need to autoclave everything what you use (unless you explicitly need it sterile). Do not touch the stuff with your bare fingers. Autoclaves can also be a souce for contamination when they are badly maintained or used to also autoclave trash. Plasticware is generally nuclease free, thanks to the production process (melted plastic is not a good place to live on).

  • If you work with RNA try to set up a separated working space to avoid contaminations by your normal work. If you use glassware, wash it with DEPC treated water and autoclave it to get it RNAse free. Use gloves! Also use a different set of chemicals for this work to avoid cross-contamination. Filter-tips can be a good idea here. Besides being careful, RNA work is no magic but solid handcraft.

  • If you do PCR try to set up a different work place seperated from your normal work. PCR is very sensitive to contaminations sitting in you pipettes from your normal work. I have seen a number of cases of false positive amplifications from normal work.

How sterile is sterile when working with nucleic acids to prevent contamination? - Biology

In addition to physical methods of microbial control, chemicals are also used to control microbial growth. A wide variety of chemicals can be used as disinfectants or antiseptics. When choosing which to use, it is important to consider the type of microbe targeted how clean the item needs to be the disinfectant’s effect on the item’s integrity its safety to animals, humans, and the environment its expense and its ease of use. This section describes the variety of chemicals used as disinfectants and antiseptics, including their mechanisms of action and common uses.


  • 1 Division of Obstetrics and Gynaecology, Faculty of Health and Medical Sciences, The University of Western Australia, Perth, WA, Australia
  • 2 Centre for Integrative Metabolomics and Computational Biology, School of Science, Edith Cowan University, Perth, WA, Australia

The human microbiome includes trillions of bacteria, many of which play a vital role in host physiology. Numerous studies have now detected bacterial DNA in first-pass meconium and amniotic fluid samples, suggesting that the human microbiome may commence in utero. However, these data have remained contentious due to underlying contamination issues. Here, we have used a previously described method for reducing contamination in microbiome workflows to determine if there is a fetal bacterial microbiome beyond the level of background contamination. We recruited 50 women undergoing non-emergency cesarean section deliveries with no evidence of intra-uterine infection and collected first-pass meconium and amniotic fluid samples. Full-length 16S rRNA gene sequencing was performed using PacBio SMRT cell technology, to allow high resolution profiling of the fetal gut and amniotic fluid bacterial microbiomes. Levels of inflammatory cytokines were measured in amniotic fluid, and levels of immunomodulatory short chain fatty acids (SCFAs) were quantified in meconium. All meconium samples and most amniotic fluid samples (36/43) contained bacterial DNA. The meconium microbiome was dominated by reads that mapped to Pelomonas puraquae. Aside from this species, the meconium microbiome was remarkably heterogeneous between patients. The amniotic fluid microbiome was more diverse and contained mainly reads that mapped to typical skin commensals, including Propionibacterium acnes and Staphylococcus spp. All meconium samples contained acetate and propionate, at ratios similar to those previously reported in infants. P. puraquae reads were inversely correlated with meconium propionate levels. Amniotic fluid cytokine levels were associated with the amniotic fluid microbiome. Our results demonstrate that bacterial DNA and SCFAs are present in utero, and have the potential to influence the developing fetal immune system.


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Allelic replacement

Escherichia coli DH5α, used for cloning, was grown in LB Lennox (BD, Difco) medium at 37°C. Mycobacterium abscessus ssp. massiliense CIP108297 was grown in Middlebrook 7H9-ADC broth (BD, Difco) supplemented with 0.05% Tween 80 or 7H11-ADC agar (BD, Difco) at 37°C. Kanamycin (Kan), streptomycin (Str) and hygromycin (Hyg) were added to final concentrations of 200, 200 and 2000 μg/ml, respectively. Homologous recombination at the trmD locus of M. abscessus ssp. massiliense CIP108297 was performed using a mycobacterial recombinase-based system in which the recombineering genes from mycobacteriophage Che9c ( 22) are expressed from the replicative plasmid pNitET-xylE-kan (a derivative of the pNitET-sacB-kan plasmid ( 23) generated in-house in which the sacB gene was replaced by the xylE colored marker) under control of an isovaleronitrile-inducible promoter. Isovaleronitrile-induced M. abscessus ssp. massiliense CIP108297 cells harboring pNitET-xylE-kan were electro-transformed with ∼300 ng of linear allelic exchange substrate consisting of the streptomycin-resistance cassette from pHP45Ω flanked by 1000 bp of DNA sequence immediately flanking the start and stop codons of trmD, and double-crossover mutants were isolated on Str-containing agar. Allelic replacement leading to the complete deletion of the trmD locus was checked by PCR using a pair of primers annealing outside the linear allelic exchange substrate.

Plasmid pMV306H was constructed by replacing the kanamycin-resistance cassette of pMV306hsp (an integrative mycobacterial expression vector allowing for the expression of genes under control of the hsp60 promoter Addgene plasmid # 26155) ( 24) by a hygromycin-resistance cassette. pMV306H::trmD was generated by cloning the PCR-amplified trmD gene from M. abscessus ssp. massiliense CIP108297 in the HindIII site of pMV306H. All primer sequences are shown in Supplementary Table S1 .

Expression and purification of full-length M. abscessus TrmD

Escherichia coli BL21 (DE3) strain containing AVA0421 plasmid with an N-His-3C Protease site-TrmD full-length insert, kindly provided by the Seattle Structural Genomics Consortium, ( 25) was grown overnight at 37°C in LB-media containing Ampicillin (100 μg/ml). This seed stage culture was used to inoculate six shake flasks containing 1 l each of 2XYT media with Ampicillin (100 μg/ml) until optical density (A600 nm) reached 0.6. The expression of recombinant construct was induced by the addition of isopropyl β- d -1-thiogalactopyranoside (IPTG) to a final concentration of 0.5 mM and further allowed to grow at 18°C for 16 h.

Isolation of cells and lysis

Cells were harvested by centrifugation at 4°C for 20 min at 5000 g and the pellet was re-suspended in buffer A (25 mM HEPES pH 7.5, 500 mM NaCl, 5% glycerol, 10 mM MgCl2, 1 mM TCEP, 20 mM imidazole). 0.1% Triton (Sigma), 10 μg/ml DNaseI, 5 mM MgCl2 and three protease inhibitor cocktail tablets (New England Biolabs) were added to the cell suspension. The cells were lysed in an Emulsiflex (Glen Creston) and clarified the lysate by centrifugation at 4°C for 40 min at 25 568 g.

Immobilized metal affinity chromatography

The clarified lysate was filtered using a 0.45 μm syringe filter and passed through a pre-equilibrated (with buffer A), 10 ml pre-packed nickel- sepharose column (HiTrap IMAC FF, GE Healthcare). The column was washed with 5 column volumes of buffer A and the bound protein was eluted as 4 × 10 ml elutes using buffer B (25 mM HEPES pH 7.5, 500 mM NaCl, 5% glycerol, 1 mM TCEP, 500 mM imidazole). The protein was analyzed on a 15% SDS-PAGE gel. Dialysis: Elutes from Hi-Trap IMAC column were pooled, added 3C Protease in the ratio of 1:50 mg (protease: protein) and subjected to dialysis against 2 l of buffer C (25 mM HEPES pH 7.5, 500 mM NaCl, 5% glycerol, 1 mM TCEP) overnight at 4°C.

Protein, after overnight dialysis and cleavage of N-His tag, was passed through a pre-equilibrated (buffer A) 5 ml HiTrap IMAC FF Nickel column (GE Healthcare).

Size exclusion chromatography

The flow through from the above column was concentrated to 3 ml using a 10 kDa centrifugal concentrator (Sartorius Stedim) and loaded onto a pre-equilibrated (with buffer D: 25 mM HEPES pH 7.5, 500 mM NaCl, 5% glycerol) 120 ml Superdex200 16/600 column (GE Healthcare). 2 ml fractions were collected and analyzed on a 15% SDS-PAGE gel. Fractions corresponding to pure TrmD protein were pooled and concentrated to 25 mg/ml, flash frozen in liquid nitrogen and stored at –80°C. Identity of the purified protein was further confirmed by MALDI mass fingerprinting.

Crystallization of apo form of full-length M. abscessus TrmD

M. abscessus TrmD apo crystals were grown in 48-well sitting drop plates (Swiss CDI) in the following condition: 0.08 mM Sodium cacodylate pH 5.8 to 6.8, 1–2 M ammonium sulphate. 24 mg/ml of the protein in storage buffer (25 mM HEPES pH 7.5, 500 mM NaCl, 5% glycerol) at drop ratio 1 μl:1 μl (protein:reservoir respectively) were set up and equilibrated against 70 μl reservoir.

Soaking of TrmD native crystals with fragments and ligands

Crystals for this experiment were grown at 19°C in 48-well sitting drop plates (Swiss CDI) in the following condition: 0.08 mM Sodium cacodylate pH 6.5 to 7.0, 1–2 M ammonium sulphate, 20 mg/ml of the protein in storage buffer (25 mM HEPES pH 7.5, 500 mM NaCl, 5% glycerol) at drop ratio 1 μl:1 μl were set up and equilibrated against 250 μl reservoir. Further, the crystals were picked and allowed to soak in a 4 μl drop containing reservoir solution and 10 mM fragments/compound (in DMSO) which was then equilibrated against 700 μl of the corresponding reservoir solution overnight at 19°C in 24-well hanging drop vapor diffusion set up.

Co-crystallization of TrmD protein with SAM/ SAH/ AW6/ AW7

2–5 mM final concentration of compound in DMSO/water was added to 20 mg/ml of TrmD protein, mixed and incubated for 2 h on ice. Crystals were grown in the following condition: 0.08 mM Sodium cacodylate pH 6.5 to 7.0, 1–2 M Ammonium sulphate or in sparse matrix screens: Wizard 1&2 (Molecular Dimensions), Wizard 3&4 (Molecular Dimensions), JCSG +Suite (Molecular Dimensions). The crystallization drops were set up at a protein to reservoir drop ratio of 0.3 μl:0.3 μl, in 96-well (MRC2) sitting drop plate, using mosquito crystallization robot (TTP labtech) and the drops were equilibrated against 70 μl of reservoir at 19°C.

X-ray data collection and processing

The TrmD apo/ligand-bound crystals were cryo-cooled in mother liquor containing 27.5% ethylene glycol. X-ray data sets were collected on I04, I02, I03, I04-1 or I24 beamlines at the Diamond Light Source in the UK, using the rotation method at wavelength of 0.979 Å, Omega start: 0°, Omega Oscillation: 0.1–0.2°, Total oscillation: 210–240°, total images: 2100–2400, Exposure time: 0.05–0.08 s. The diffraction images were processed using AutoPROC ( 26), utilizing XDS ( 27) for indexing, integration, followed by POINTLESS ( 28), AIMLESS ( 29) and TRUNCATE ( 30) programs from CCP4 Suite ( 31) for data reduction, scaling and calculation of structure factor amplitudes and intensity statistics. All TrmD crystals belonged to space group P212121 and consisted of two protomers in the asymmetric unit.

Structure solution and refinement

The M. abscessus TrmD Apo structure was solved by molecular replacement using PHASER ( 32) with the atomic coordinates of M. abscessus TrmD at 1.7 Å (PDB entry: 3QUV Seattle Structural Genomics Consortium for Infectious Diseases) as search model and TrmD ligand bound structures were solved by molecular replacement with the atomic coordinates of the solved M. abscessus TrmD Apo structure (PDB entry: 6NVR) as search model. Structure refinement was carried out using REFMAC ( 33) and PHENIX ( 34).

The models obtained were manually re-built using COOT interactive graphics program ( 35) and electron density maps were calculated with 2|Fo| – |Fc| and |Fo| – |Fc| coefficients. Positions of ligands and water molecules were located in difference electron density maps and OMIT difference maps |mFo − DFc| ( 36) were calculated and analysed to further verify positions of fragments and ligands.

Differential scanning fluorimetry (DSF)

DSF were carried out in a 96-well format with each well containing 25 μl of reaction mixture of 10 μM TrmD protein in buffer (50 mM HEPES pH 7.5, 500 mM NaCl, 5% glycerol), 5 mM compound, 5% DMSO and 5× Sypro orange dye. Appropriate positive (Protein, DMSO and SAM) and negative (Protein, DMSO only) controls were also included. The measurements were performed in a Biorad-CFX connect thermal cycler using the following program: 25°C for 10 min followed by a linear increment of 0.5°C every 30 s to reach a final temperature of 95°C. The results were analyzed using Microsoft excel.

Isothermal titration calorimetry (ITC)

ITC experiments to quantify binding of ligands to TrmD were done as described in ( 37) using Malvern MicroCal iTC200 or Auto-iTC200 systems at 25°C. Titrations consisted of an initial injection (0.2 μl), discarded during data processing, followed by either 19 (2 μl) or 39 (1 μl) injections separated by intervals of 60–150 s duration. Protein was dialysed overnight at 4°C in storage buffer (M. abscessus TrmD: 50 mM HEPES pH 7.5, 500 mM NaCl, 5% glycerol M. tuberculosis TrmD: 25 mM HEPES pH 7.5, 500 mM NaCl). Sample cell and syringe solutions were prepared using the same storage buffer, with a final DMSO concentration of 2–10% according to ligand solubility in the buffer. TrmD concentrations of either 33 or 100 μM were used, with ligand to protein concentration ratios ranging from 10 to 20:1. Control titrations without protein were also performed and subtracted from ligand to protein titrations. Titrations were fitted with Origin software (OriginLab, Northampton, MA, USA), using a one-site binding model with N fixed to 1 only for weakly binding ligands. Titrations were typically performed once (n = 1), with multiple isotherms obtained (n > 1) for key compounds of interest. Kd values are reported to two significant figures. Error provided by Origin software due to model fit is reported when n = 1, whereas standard deviation is reported when n > 1.

Chemical synthesis of compounds

The compounds AW1-7 were synthesised according to the procedures described in the supporting information. A more in-depth discussion around the medicinal chemistry strategy for fragment merging and lead compound development is described in a corresponding publication by Whitehouse et al. ( 37). The compounds AW1-7 are listed in this publication as follows: AW1 (Compound 23), AW2 (Compound 24f), AW3 (Compound 26f), AW4 (Compound 28), AW5 (Compound 29a), AW6 (Compound 31a) and AW7 (Compound 29d).

Biochemical activity assays

Assays for quantifying TrmD methylation reactions were carried out in 20 μl reactions consisting of 6.25 μM SAM, 0.1 μM TrmD and 6.25 μM tRNA Pro UGG in the presence of 0–500 μM compounds in serial dilutions using assay buffer containing 50 mM Tris–HCl pH 7.5, 10 mM MgCl2, 24 mM NH4Cl, 5% DMSO and 1 mM DTT in nuclease free water. tRNA sequences were identified from the M. abscessus genome sequence using tRNAscan-SE algorithm, ( 38, 39). The substrate M. abscessus tRNA Pro UGG for the assay, having the sequence 5′-CGGGGUGUAGCGCAGCUUGGUAGCGCAUCCGCUUUGGGAGCGGAGGGUCGCAGGUUCAAAUCCUGUCACCCCGA-3′, was purchased commercially from Integrated DNA technologies (USA). The reactions were carried out for 1 h at room temperature followed by addition of 20 mM EDTA to stop the reactions. Each of the 20 μl samples were diluted ten-fold with the UPLC mobile phase solvent A (0.1% formic acid in water), centrifuged for 10 min at 13 000 g, to remove any precipitates, and the supernatant was aliquoted into 96-well plates. 40 μl samples were then injected into Acquity UPLC (Waters) T3 1.8 μM column and eluted using a gradient elution consisting of Mobile Phase A: 0.1% formic acid in water and mobile phase B: 0.1% formic acid in 100% methanol for 4 min. The absorbance was monitored using a photodiode array (PDA) detector (Waters) at wavelength range of ƛ: 220–500 nm. All reactions were carried out in triplicate. The blank corrected data were analyzed using Microsoft excel and non-linear regression analysis for IC50 determination were done using GraphPad prism version 7.00, GraphPad Software, La Jolla, CA, USA.

Mycobacterial strains used and MIC measurements

Mycobacterium abscessus ssp. abscessus (ATCC 19977) transformed with pmv310 plasmid expressing Lux ABDCE operon, grown in Middlebrook 7H9 broth supplemented with ADC (Sigma, UK). All the other NTM strains are clinical isolates. Minimum inhibitory concentrations (MIC) were determined for mycobacteria according to the Clinical and Laboratory Standards Institute (CLSI) method M07-A9. Briefly, mycobacteria were grown to optical density (A600 nm) of 0.2–0.3 in liquid culture and 1 × 10 5 bacteria were added to each well of 96-well plates containing serial dilutions of compound (400, 200, 100, 50, 25, 12.5, 6.3, 3.1, 1.6, 0.8, 0.4, 0 μM), in triplicate wells per condition, and incubated at 37°C until growth was seen in the control wells. MIC measurements using M. tuberculosis H37Rv were performed as reported in ( 37). M. tuberculosis H37Rv was grown in Middlebrook 7H9 base containing 14 mg/l dipalmitoyl phosphatidylcholine (DPPC), 0.81 g/l NaCl, 0.3 g/l casitone, and 0.05% Tyloxapol. H37Rv was grown and diluted to a similar inoculum size as mentioned above prior to exposure to serial dilutions of compounds (starting at 100 μM), and the plates were incubated at 37°C for 2 weeks. The MIC value was determined as the last well which showed no bacterial growth.

CRISPR–dCas9 knockdown in M. abscessus

The dCas9 encoding plasmid (pTetInt-dcas9-Km) and the second vector containing the sgRNA cassette (pGRNAz) were derived from the tetracycline inducible CRISPr Interference system of Choudhary et al. ( 40) and optimized for M. abscessus ATCC19977. The 20 nucleotides guides targeting yidC(MAB_4953c) and trmD(MAB_3226c) were annealed and cloned in between sphI and aclI of the pGRNAz. As a control the pGRNAz was left empty. The CRISPr-I containing strains were cultivated in Middlebrook 7H9 broth supplemented with 1× ADC, 0.05% tween80 and 0.5% glycerol, hygromycin 1 mg/ml and zeocin 300 μg/ml. The cultures were inoculated at 10^6CFU/ml from an exponentially grown pre-cultures. The AW7 compound and doxycycline were added or not at a concentration of 25μM and 1.5625, 6.25, 25 or 100 ng/ml respectively. The OD600 was measured and the CFU counted at 72 h.

Macrophage infection study

Blood samples were donated by healthy volunteers who had undertaken informed consent in accordance with local Research Ethics Committee approval. Peripheral blood mononuclear cells were isolated from citrated peripheral blood samples by density gradient separation using Lympholyte (Cedarlane Labs), and subsequent CD14 + positive selection using the MACS Miltenyi Biotec Human CD14 microbead protocol (Miltenyi Biotec). CD14 + cells were differentiated into macrophages using recombinant human granulocyte-macrophage colony-stimulating factor (200 ng/ml GM-CSF) and recombinant human interferon gamma (50 ng/ml IFNγ) (Peprotech) in standard tissue culture DMEM media containing fetal calf serum, penicillin and streptomycin. Following removal of antibiotics, macrophages were infected at a multiplicity of infection of 10:1 with M. abscessus 19977 for 2 h, washed in sterile phosphate buffered saline, and then incubated in DMEM media with FCS and 25 μM of compound for 24 and 48 h. At the given time points, supernatant was saved for cell cytotoxicity studies, and M. abscessus survival within the macrophages calculated by macrophage lysis in sterile water, and colony forming unit calculation on Columbia Blood Agar plates (VWR BDH).


Lactate dehydrogenase (LDH) was measured as a biomarker for cellular cytotoxicity using the Pierce LDH Cytotoxicity Assay Kit. Cell supernatant was measured at 2, 24 and 48 h post-infection according to the kit protocol.

Nude mouse derived M. leprae

Mycobacterium leprae (isolate Thai-53) was maintained in serial passage in the foot pads of athymic nude mice (Envigo, USA). Mice were inoculated in the plantar surface of both hind feet with 5 × 10 7 fresh, viable nude mice derived M. leprae. When the mouse foot pads became moderately enlarged (at ∼5–6 months), they were harvested for intracellular M. leprae as described previously ( 41), washed by centrifugation, re-suspended in medium, enumerated by direct count of acid fast bacilli according to Shepard's method ( 42), held at 4°C pending quality control tests for contamination and viability ( 41). Freshly harvested bacilli were always employed in experiments within 24 h of harvest.

M. leprae axenic culture

Freshly harvested nude mouse foot pad derived M. leprae were suspended in modified 7H12 medium, AW7 was added at different concentrations (100–6.25 μM) and were incubated for 7 days at 33°C. Media only and rifampin (Sigma, USA) at 2.4 μM were used as negative and positive controls. Following incubation aliquots of AW7 treated and control M. leprae were processed for radiorespirometry (RR) as described previously ( 43).

M. leprae macrophage culture

Bone marrow cells were obtained aseptically from both femurs of female BALB/c mice and cultured on plastic cover slips in Dulbecco modified Eagle's medium (DMEM, Life Technologies, USA) supplemented with 10% (v/v) fetal calf serum (Life Technologies), 25 mM/l HEPES (Sigma, USA), 2 mM/l glutamine (Sigma, USA), 50 μg/ml ampicillin (Sigma, USA) and 10 ng/ml of recombinant murine macrophages colony stimulating factor (R&D Systems, USA) for 6–7 days at 37°C and 5% CO2. The cells were infected with freshly harvested nude mice foot pad derived live M. leprae at a multiplicity of infection (MOI) of 20:1 overnight at 33°C and then washed to remove extracellular bacteria. AW7 was added at different concentrations (100–6.25 μM) and the cells were incubated for 7 days at 33°C. Media only and rifampicin at 2.4 μM were used as negative and positive controls. AW7 treated and control cells were lysed with sodium dodecyl sulfate (SDS, 0.1% w/v, Sigma, USA) and the intracellular M. leprae processed for radiorespirometry ( 44).


Metabolism of a suspension of M. leprae was measured by evaluating the oxidation of 14 C-palmitic acid to 14 CO2 by radiorespirometry as described previously ( 45). Levels of captured 14 CO2 is proportional to the rate of 14 C-palmitic acid oxidation and used as an indicator of M. leprae viability. In the present study the seventh day cumulative counts per minute (CPM) were recorded and percentage inhibition of metabolism determined as compared to no drug control. Statistical significance between treatment groups and no drug control were determined by Student's t-test and P < 0.05 is considered as significant.

Uric acid promotes an acute inflammatory response to sterile cell death in mice

1 Department of Pathology, University of Massachusetts Medical School, Worcester, Massachusetts, USA. 2 Department of Biochemical Science and Technology, National Taiwan University, Taipei, Taiwan.

Address correspondence to: Kenneth L. Rock, Department of Pathology, University of Massachusetts Medical School, 55 Lake Ave. North, Worcester, Massachusetts 01655, USA. Phone: 508.856.2521 Fax: 508.856.1094 E-mail: [email protected]

1 Department of Pathology, University of Massachusetts Medical School, Worcester, Massachusetts, USA. 2 Department of Biochemical Science and Technology, National Taiwan University, Taipei, Taiwan.

Address correspondence to: Kenneth L. Rock, Department of Pathology, University of Massachusetts Medical School, 55 Lake Ave. North, Worcester, Massachusetts 01655, USA. Phone: 508.856.2521 Fax: 508.856.1094 E-mail: [email protected]

1 Department of Pathology, University of Massachusetts Medical School, Worcester, Massachusetts, USA. 2 Department of Biochemical Science and Technology, National Taiwan University, Taipei, Taiwan.

Address correspondence to: Kenneth L. Rock, Department of Pathology, University of Massachusetts Medical School, 55 Lake Ave. North, Worcester, Massachusetts 01655, USA. Phone: 508.856.2521 Fax: 508.856.1094 E-mail: [email protected]

Find articles by Ontiveros, F. in: JCI | PubMed | Google Scholar

1 Department of Pathology, University of Massachusetts Medical School, Worcester, Massachusetts, USA. 2 Department of Biochemical Science and Technology, National Taiwan University, Taipei, Taiwan.

Address correspondence to: Kenneth L. Rock, Department of Pathology, University of Massachusetts Medical School, 55 Lake Ave. North, Worcester, Massachusetts 01655, USA. Phone: 508.856.2521 Fax: 508.856.1094 E-mail: [email protected]

Related article:

Caught red-handed: uric acid is an agent of inflammation

Caught red-handed: uric acid is an agent of inflammation


Inflammation occurs in response to both pathogenic insult and tissue damage under sterile conditions, with the latter contributing to the pathogenesis of many diseases. Although several endogenous substances, including uric acid, have been suggested to alert the body to danger and to stimulate inflammation, little is known about their contribution to such responses in vivo. In this issue of the JCI, Kono et al. use newly generated mice with reduced levels of uric acid to investigate its role as an endogenous signal of tissue damage in inflammatory responses to hepatic injury. They find that uric acid is released from dying tissues and induces inflammation to cell death but is not involved in the response to microbial molecules or sterile irritant particles. I believe this to be the first report of an endogenous danger signal acting as a physiological regulator of inflammation.


Necrosis stimulates inflammation, and this response is medically relevant because it contributes to the pathogenesis of a number of diseases. It is thought that necrosis stimulates inflammation because dying cells release proinflammatory molecules that are recognized by the immune system. However, relatively little is known about the molecular identity of these molecules and their contribution to responses in vivo. Here, we investigated the role of uric acid in the inflammatory response to necrotic cells in mice. We found that dead cells not only released intracellular stores of uric acid but also produced it in large amounts postmortem as nucleic acids were degraded. Using newly developed Tg mice that have reduced levels of uric acid either intracellularly and/or extracellularly, we found that uric acid depletion substantially reduces the cell death–induced inflammatory response. Similar results were obtained with pharmacological treatments that reduced uric acid levels either by blocking its synthesis or hydrolyzing it in the extracellular fluids. Importantly, uric acid depletion selectively inhibited the inflammatory response to dying cells but not to microbial molecules or sterile irritant particles. Collectively, our data identify uric acid as a proinflammatory molecule released from dying cells that contributes significantly to the cell death–induced inflammatory responses in vivo.

When cells undergo necrosis in vivo, they trigger an acute inflammatory response ( 1 ). Local blood flow is increased, venules leak protein-rich fluid, and leukocytes extravasate from the blood into the tissue. Neutrophils are the first cells recruited into the site of cell death, and this is followed later by an influx of monocytes. These events occur wherever significant necrosis occurs and are so reproducible that the progression of the inflammatory response is used forensically to date the time of tissue injury ( 2 ).

Why should cell death stimulate an inflammatory response? Necrosis is not typically a normal physiological event but rather occurs as a consequence of some pathological process that damages cells as such, necrosis is a harbinger of a potential threat to the host ( 3 ). Presumably because of this, the innate immune system is rapidly mobilized to deliver the soluble and cellular defenses to a site of injury. Once deployed, the defenses will engage and attempt to neutralize or wall off any injurious agents that are present. They also help to clear dead cells and debris and stimulate mechanisms to repair tissue damage. On the other hand, recruiting the innate defenses comes at a price ( 4 – 6 ). The delivery of the immune effector mechanisms is imprecise, and as a result, normal healthy cells may be caught in the line of fire and damaged. Hydrolytic enzymes and highly reactive chemical compounds, such as oxygen radicals, leak from living and dying leukocytes, and these molecules inflict damage on cells in the environment. In infections, this process is often short lived and the resulting collateral damage is a small price to pay to contain a potentially life-threatening condition. However, in situations of sterile cell death, and when this process is chronic, the cost-benefit ratio is less favorable and can lead to disease. Because of these positive and negative effects, it is important to understand the mechanisms by which cell death leads to inflammation ( 7 , 8 ).

How does cell death actually provoke an inflammatory response? It is thought that cells contain immune stimulatory molecules (often referred to as damage-associated molecular patterns [DAMPs]) that are not exposed in living cells, but are released upon necrosis ( 9 – 11 ). The innate immune system has evolved the capacity to recognize and then respond to the presence of DAMPs. There may be different classes of DAMPs that act on different cellular targets and have different biological effects. For example, some DAMPs may trigger inflammation (referred to herein as proinflammatory DAMPs) ( 12 ) and others may function as adjuvants to promote adaptive immune responses ( 13 ), while yet others may affect other processes such as tumor behavior ( 14 ).

The molecular identity of the proinflammatory DAMPs is very poorly understood. A few potential candidate molecules have been recently described, including molecules such as HMGB1 ( 12 ), SAP130 ( 15 ), myosin heavy chain ( 16 ), and fragments of extracellular matrix components (generated from hydrolases released from dead cells) ( 17 , 18 ). However while these molecules have proinflammatory potential, their actual role in cell death–induced inflammation in vivo is for the most part unclear. For example, while HMGB1 can induce inflammation ( 12 ), necrotic mutant cells lacking HMGB1 are as proinflammatory as WT cells ( 19 ) whether this is because HMGB1 is not truly a proinflammatory DAMP in vivo or redundancy in proinflammatory DAMPs makes its presence nonessential is not clear. Determining the molecular identity of DAMPs and their contribution to inflammation in vivo is important because these molecules could be targets for therapeutic intervention to prevent or treat diseases caused by cell death–induced inflammation.

We had previously found that uric acid is released from dying cells, where it then functions as an adjuvant that promotes the generation of adaptive immune response ( 13 , 20 ). It is thought that the active form of this molecule is monosodium urate, which acts to promote immune responses by stimulating dendritic cells. Independent of this and in other settings, monosodium urate crystals can trigger inflammation, and when this occurs spontaneously in hyperuricemic patients, it causes the inflammatory disease of gout ( 21 ). These findings have led us to hypothesize that uric acid released from dying cells might function as a proinflammatory DAMP that contributes to cell death–induced inflammation. The studies in this report were designed to test this hypothesis.

Uricase Tg mouse models. To analyze the roles of uric acid in vivo, it would be useful to have animals that are deficient in this molecule. It is possible to lower the levels of uric acid in animals using pharmacological approaches. One approach has been to block its synthesis. Uric acid is generated from xanthine by xanthine oxidase, and this reaction can be inhibited by the uric acid analogue allopurinol ( 22 ). A limitation of this approach is that it only partially reduces but does not eliminate uric acid from cells and body fluids. Another approach is to deplete uric acid by administering uricase, an enzyme, which oxidizes uric acid into allantoin and water. This treatment can markedly lower uric acid levels but has limitations in its pharmacokinetics, including slow distribution into interstitial fluids and rapid clearance from the blood also, currently available sources of this enzyme are immunogenic in mice and elicit neutralizing antibodies, precluding long term use in vivo ( 23 ). Because of these issues, it would be attractive to have genetic models that are stably deficient in uric acid that could be used in addition to the pharmacological treatments. Unfortunately, mice that are genetically deficient in xanthine oxidase, and therefore unable to produce uric acid, are runted at birth and die after several weeks ( 24 ).

We decided to pursue a different approach to develop uric acid–deficient mice. We hypothesized that the expression of murine uricase as a transgene might be able to chronically reduce uric acid levels in tissues and overcome some of the limitations of the other available approaches. Uricase is only expressed in the liver of mice. We cloned this gene from mouse liver and used it to produce 2 different constructs. In one construct, the uricase cDNA was fused with a signal sequence from the adenovirus E3-gp19K gene so that when expressed, the uricase protein would be secreted into the extracellular fluids (Figure 1A and ref. 25 ). The idea here was that the secreted uricase would deplete uric acid from the interstitial fluids of tissues. In another construct, the uricase cDNA was targeted to peroxisomes using its endogenous targeting sequence (Figure 1A this is referred to herein as the intracellular uricase construct). The idea behind this construct was that the transgene should reduce intracellular pools of uric acid and, after cells die and release the enzyme, also deplete uric acid from the environment of dead cells. Thus these 2 Tg animals were designed to lower uric acid levels around dying cells but to do so in different ways. Both the secreted and intracellular uricase constructs were placed under the control of the β actin promoter so that they would be expressed and exert their effects broadly in most tissues ( 26 ). These uricase constructs were injected into the nuclei of fertilized C57BL/6 eggs, and several independent Tg founders were generated. The animals were viable, fertile, and without any obvious phenotypic abnormalities. A third construct was also generated wherein the peroxisomal targeting sequence was deleted so that the uricase would be expressed in the cytosol however, cells did not tolerate stable expression of this cDNA, so it was not used in our studies (data not shown).

Generation and characterization of uricase Tg mice. (A) Construct of uricase transgenes. Secreted uricase (ssUOX) was generated by N-terminal addition of a signal sequence for secretion derived from adenovirus gp19K (gp19K-ss). The unmodified intracellular uricase (intUOX) has a C-terminal peroxisome targeting signal sequence (PTS). (B) Western blot of uricase and α-tubulin (loading control) in organs of Tg or WT mice. (C) Uricase activity in organs and serum. WT C57BL/6 mice were injected with 9 μg of i.p. and 9 μg of i.v. rasburicase where indicated. Organs were harvested from untreated WT, uricase Tg mice, or WT mice 18 hours after rasburicase injection, and lysates were prepared. Twenty μl of lysate form various organs was added to 1 ml of uric acid solution (OD292 = 1.0) and incubated at 37°C for indicated periods of time. The uricase activity was measured by the decrease of OD292. (D) Amount of uric acid in peritoneal cavity in WT and uricase Tg mice (n = 6). (E) Plasma concentration of uric acid in WT and uricase Tg mice. Samples were drawn and immediately chilled on ice to prevent uricase from oxidizing uric acid ex vivo (n = 13–19). (F) Total neutrophil numbers in the peritoneal cavity after 15 hours after i.p. injection of 2 mg of monosodium urate crystal. n = 6 (PBS) n = 15 (WT) n = 8 (ssUOX). **P < 0.01 *P < 0.05 versus WT in (DF).

We analyzed the expression of uricase by Western blot in serum and tissue lysates from WT and Tg animals. Uricase was undetectable in the serum of WT mice and those Tg for the intracellular construct, as expected (Figure 1B). In contrast, uricase was present in the serum of mice Tg for the signal sequence–uricase construct, indicating that the transgene was secreted, as expected (Figure 1B). Presumably most of the enzyme in the serum originates from the interstitial fluids (delivered after drainage through lymphatics) and possibly also from cells in the blood and tissues with sinusoidal architecture (e.g., the spleen, liver, and bone marrow). In WT animals, uricase was undetectable in all tissues except for the liver, as expected (Figure 1B). In contrast, uricase was detectable in all tissues of both Tg animals, including in the liver, where it was present in elevated amounts compared with WT mice (Figure 1B). Therefore, the Tg uricase constructs were expressed with the expected distribution.

To verify that the Tg uricase was active, we also performed enzyme assays on serum and tissue lysates. Uricase activity was only detected in the serum of the secreted uricase Tgs and not the intracellular ones or WT animals. The lysates from all organs examined from both Tgs contained active uricase (Figure 1C), while all tissues from WT mice except liver lacked this activity. The content of the secreted uricase varied between different tissue lysates, presumably due to differences in the amount of this enzyme in the ER, exocytic pathway, and possibly extracellular fluid. Interestingly, lysates of most tissues from animals injected i.v. 18 hours earlier with recombinant uricase had little or no uricase activity, presumably because the enzyme remained intravascular and/or was cleared the exception was the lung (Figure 1C). These results indicate that recombinant intracellular and secreted forms of uricase are enzymatically active, both in cells and, when secreted, in the extracellular environment and serum of mice.

Uric acid levels in uricase Tg mice. We next examined the effects of the expressed uricase on uric acid levels in the various Tg mice. In the mice expressing secreted uricase, we predicted that there would be a reduction in uric acid levels in the interstitial fluids. We evaluated this prediction in the resident peritoneal fluid sampled by lavage. As predicted, uric acid levels in peritoneal fluid of the secreted uricase Tg mice were significantly reduced compared with WT animals, although the reduction was not complete (

60% Figure 1D). We also measured uric acid levels in freshly isolated serum and found that they were reduced by approximately 30% in the secreted uricase Tg animals although this reduction was more modest than in the peritoneum, it was significant (P < 0.05 Figure 1E). The greater reduction in uric acid in the tissue fluids compared with the serum is likely to reflect the direct secretion of uricase into the tissue fluids, where because of the small volume of fluid, the enzyme is presumably at higher concentration than after it is resorbed and distributed throughout the serum. To determine whether the secreted uricase was sufficient to inhibit inflammatory responses to uric acid, we injected monosodium urate crystal i.p into control or ssUOX Tg mice and found that the resulting inflammatory responses were significantly inhibited by the secreted uricase (Figure 1F).

Upon incubating the serum of WT animals, ex vivo uric acid levels were stable (Figure 2A). There was a slow and modest reduction in uric acid levels upon incubation of serum from the intracellular Tg mice, indicating the presence of a small amount of uricase (Figure 2A). In contrast after incubation of serum from the secreted uricase Tg mice, uric acid levels rapidly fell to undetectable levels, reflecting the activity of the uricase present in the serum (Figure 2A). The fact that uric acid was still present at all in the presence of active uricase in the serum indicates that in vivo, the production of uric acid exceeds its removal. Since uric acid levels are regulated in vivo by cellular production as well as adsorption from the gut and readsorption in the kidney ( 27 ), it is possible that in the Tg mice, there have been compensatory increases in these processes in response to the loss of uric acid however, this issue has not been investigated. There was no increase in uric acid in homogenates of any tissue in ssUOX Tg mice, and rather some decrease was found in several organs. In any case, these data indicate that the secreted uricase Tg mice have a significant reduction in uric acid levels in extracellular fluids and serum.

Generation of uric acid after cell death. (A) Change of uric acid concentration in serum of WT and uricase Tg mice incubated ex vivo. Serum from uricase Tg mice was collected after being allowed to clot at room temperature and subsequently incubated at 37°C for the indicated times uric acid concentration was measured (n = 32–36). (B) Generation of uric acid after cell death. Tissue lysates obtained from the indicated organs from uricase Tg mice or WT littermates were incubated at 37°C for indicated times. Uric acid content was normalized with protein concentration. (C) Postmortem uric acid is generated by xanthine oxidase. Tissue lysates were incubated at 37°C for indicated times in the presence of allopurinol (128 mg/l) or uricase (0.5 mg/ml). The uric acid concentrations in the tissue lysate are shown.

In mice Tg for the intracellular uricase construct, we examined the intracellular levels of uric acid. In the majority of Tg tissues, we found a partial but significant reduction (30%–70%) in intracellular levels of uric acid compared with WT cells (Figure 2B and Supplemental Figure 1 supplemental material available online with this article doi: 10.1172/JCI40124DS1). Interestingly, when the dead cells were cultured at 37°C, as would occur when cells die in vivo, we observed a marked increase in uric acid levels in WT cells. This increase in synthesis was blocked by inhibitors of xanthine oxidase, indicating that purines were being metabolized into uric acid postmortem (Figure 2C). Remarkably, this postmortem increase in uric acid was suppressed in both intracellular and secretable uricase Tg mice. These data indicate that the intracellular and secretable recombinant uricase is released from dead cells and eliminates the endogenous and newly produced uric acid in the local environment. Serum levels of uric acid were not reduced in the intracellular uricase Tgs (Figure 1E), and this was not surprising, given that a substantial amount of uric acid originates from the diet (absorbed from the gut) and that there was only a partial reduction in intracellular stores of uric acid in the Tgs. Moreover, there is little of the intracellular uricase released into the serum (Figure 1B and Figure 2A).

In summary, the 2 Tg animals have different distributions of uricase and correspondingly partial but significant reductions in uric acid in different body compartments. The secreted uricase Tg has uricase and reduced uric acid in the extracellular fluids. The intracellular uricase Tg animals have uricase intracellularly that reduces intracellular, but not extracellular, pools of uric acid. In addition, the Tg uricase is released in active form from dead cells and then eliminates uric acid in and around the dead cells. Thus these 2 different transgenes will reduce uric acid around dying cells but do so in somewhat different ways.

Effect of uric acid depletion on inflammation to cell death in the liver. To examine the role of uric acid in cell death–induced inflammation, we used a well-established liver injury model in which mice were treated with a toxic dose of acetaminophen ( 19 ). This drug is metabolized in the liver to N-acetyl-p-benzoquinone imine, which causes necrosis in the centrilobular regions of this organ. The resulting sterile cell death stimulates a robust neutrophilic inflammatory response, which can be quantified by measuring the content of a neutrophil-specific enzyme myeloperoxidase (MPO) in the liver ( 19 ). The extent of liver injury can be quantified by measuring the amount of liver cytosolic enzymes (alanine aminotransferase [ALT]) released and present in the serum.

The 2 uricase Tg mouse models and WT mice were treated with a single toxic dose of acetaminophen. The resulting inflammatory response to cell death in the liver was quantified and found to be very different in the WT versus uricase Tg mice. The injured liver in WT animals developed robust neutrophilic inflammation, while the inflammation in the injured livers of Tg mice was markedly reduced (Figure 3A). The reduction in the inflammatory response was seen both in the intracellular and secreted uricase Tg mice, with a somewhat larger inhibition with the intracellular construct (Figure 3A). The inflammatory cell infiltrate in the treated livers was further characterized by dissociating the liver into a single-cell separation followed by immunofluorescent staining and flow cytometry (Supplemental Figure 2). Eighteen hours after injury, the predominant inflammatory cells were neutrophils (as expected) and their numbers were significantly reduced in both uricase Tgs. There were many fewer monocytes, macrophages, T cells, or B cells at this time point after injury, and their numbers were not significantly reduced in either of the uricase Tgs (Supplemental Figure 2). Immunohistochemical detection of neutrophils by staining for esterase activity similarly revealed infiltrating neutrophils and a reduction in their numbers in both of the uricase Tg mice (Figure 3, B and C). In addition to measuring MPO in the liver, we assayed the levels of proinflammatory cytokines in the control and treated animals. We could detect elevated levels of IL-1βMIP2 and KC of treated control animals, and the levels of these proinflammatory mediators were significantly reduced in the serum and/or liver of the UOX-Tg mice (Supplemental Figure 3).

Reduced neutrophil recruitment to liver injury in uricase Tg mice. Liver tissue MPO activity (A), number of esterase-positive cells (neutrophils) (B), histology of sections stained for esterase activity (C), serum ALT activity (D), quantification of necrotic area (E), and histology of liver sections stained with H&E (F) of WT and uricase Tg mice 18 hours after challenge with 300 mg/kg acetaminophen. Total numbers of mice used in A and D from 4 independent experiments were n = 6 (PBS control) n = 26 (WT) n = 24 (ssUOX) and n = 23 (intUOX). Total numbers of mice used in B and E were n = 6 (PBS control) n = 8 (WT) n = 5 (ssUOX) and n = 5 (intUOX). Means and SEM values are shown. **P < 0.01 *P < 0.05. NS, not significant versus control WT C57BL/6. (C) Representative images of esterase staining used in the analysis of B. Arrowheads indicate the esterase-positive cells. Scale bars: 25 mm. (F) Representative images of H&E staining used in the analysis of E. Scale bars: 250 mm.

In contrast, ALT enzyme levels rose in the serum of both WT and Tg mice to levels that were not significantly different, indicating that the amount of drug-induced hepatocellular injury was similar and not affected by uricase expression (Figure 3D). This was further confirmed by histopathological examination of liver sections both WT and both uricase Tg mice showed similar levels of centrilobular necrosis (Figure 3, E and F). At this early time point, there was no evidence of fibrosis or vascular damage, as expected. Therefore, the expression of uricase was inhibiting the inflammatory response to hepatocyte injury and doing so whether the transgene was secreted or intracellular. It should be noted we and others have previously reported that liver cell damage can be reduced by blocking inflammation ( 19 , 28 ) and this protective effect was not seen in these experiments this is presumably because inflammation needs to be inhibited almost completely to see this effect.

These results suggest that uric acid is released from (and based on the data in Figure 2B, potentially further generated by) dying hepatocytes and that the UOX reduces the levels of this molecule in the injured liver. To further investigate this issue, we measured the content of uric acid in the livers of controls and acetaminophen-treated mice. Uric acid levels were significantly elevated in the injured liver of control mice, and these levels were reduced in both the ssUOX and intracellular UOX Tg mice (Supplemental Figure 4).

We also examined the effect in this model of reducing uric acid concentrations acutely through injection of recombinant uricase. This treatment also did not reduce the amount of liver injury caused by acetaminophen injection, as assessed by the release of ALT into the serum (Figure 4B). However, the recombinant uricase did reduce uric acid levels in the liver (Supplemental Figure 4) and cause a partial but significant reduction in liver inflammation after injury (Figure 4A). The degree of inhibition of inflammation was less than that seen in the uricase Tg animals, presumably because the injected uricase is less bioavailable at the site of injury. The fact that qualitatively similar results were obtained when uric acid levels were reduced by the expression of the transgenes or acute injection of uricase argues against the possibility that chronic uric acid depletion in the Tgs might somehow be leading to adaptations that indirectly affected the inflammatory response.

Reduced neutrophil recruitment to liver injury in mice treated with rasburicase. Liver tissue MPO activity (A) and serum ALT activity (B) of control and rasburicase-treated mice 18 hours after challenge with 300 mg/kg acetaminophen. n = 6 (PBS) n = 20 (APAP) n = 18 (APAP + rasburicase). Means and SEM values are for combined data from 4 independent experiments. **P < 0.01, versus control APAP alone group. NS, not significant versus APAP alone.

In another set of experiments, we investigated the effect on inflammation to tissue injury of reducing uric acid levels using allopurinol to block its synthesis. We confirmed that allopurinol treatment lowered intracellular concentrations of uric acid in the liver (Figure 5A) and found that it did not significantly reduce its levels in serum in this time period (not shown). Although this treatment similarly did not significantly reduce the amount of liver injury caused by acetaminophen injection, it did inhibit the ensuing inflammatory response (Figure 5, B and C). This result is important because it confirms the results obtained with Tg and injected uricase and does so by lowering uric acid though a completely different mechanism. This argues strongly that the reduced inflammation to cell death is due to uric acid depletion and not some other pleiotropic effect of the treatments.

Reduced neutrophil recruitment to liver injury by allopurinol treatment. (A) Uric acid concentration in liver lysate treated with 1 week of i.p. injections of allopurinol (10 mg/kg/d). Liver tissue MPO activity (B) and serum ALT activity (C) of control and allopurinol-treated mice 18 hours after challenge with 300 mg/kg acetaminophen. n = 6 (PBS) n = 17 (APAP) n = 17 (APAP + allopurinol). Means and SEM values are for combined data from 3 independent experiments. *P < 0.05, versus control APAP alone group. NS, not significant versus APAP alone.

Effect of uric acid depletion on inflammation to other necrotic cells in the peritoneal cavity. To determine whether uric acid was playing a role in death-induced inflammation to other cell types and in another anatomic location, we evaluated the effect of depleting uric acid on inflammatory responses to dead cells injected into the peritoneum. Injection of necrotic EL4 (a syngeneic T cell lymphoma) into the peritoneum elicits strong neutrophilic inflammation (Figure 6A), as previously reported ( 19 ). The number of cells (3 × 10 7 ) injected was chosen to obtain robust responses and are relevant to in vivo situations where even much larger numbers of cells die under pathological conditions (e.g., myocardial infarct or toxic damage). This inflammatory response is partially but significantly reduced when uric acid is depleted by injection of recombinant uricase (Figure 6A). We similarly found that the inflammatory response to dead cells injected i.p. was reduced in the ssUOX Tg mice (Supplemental Figure 5).

Reduced neutrophil recruitment in the peritoneal cavity in response to rasburicase-treated necrotic cells or lung from uricase Tg mice. Total neutrophil or monocyte numbers in the peritoneal cavity 15 hours after i.p. challenge with necrotic EL4 cells (A and B) or lung homogenate from WT C57BL/6 (C and D) with or without 18 μg of rasburicase. (E and F) Lung homogenate from WT or intracellular uricase Tg mice was injected i.p. into WT C57BL/6. Neutrophil or monocyte numbers in PEC were determined by counting the Ly-6G + 7/4 + or Ly-6G – 7/4 + cells in 100 μl of peritoneal lavage, respectively. Negative control: C57BL/6 mice challenged with PBS. Uric acid concentrations of EL4 cell suspension and WT lung homogenate injected were 5.4 mg/dl and 7.5 mg/dl, respectively. Means and SEM values are for combined data from 3 or more independent experiments. Number of mice used in (A and B): n = 8 (PBS) n = 15 (EL4) n = 16 (EL4 + rasburicase) (C and D): n = 10 (PBS) n = 24 (lung) n = 23 (lung + rasburicase) (E and F): n = 4 (PBS) n = 10 (WT-lung) n = 10 (intUOX Tg-lung). **P < 0.01 NS, not significant versus control necrotic EL4 without rasburicase (A and B), lung homogenate without rasburicase (C and D), or WT-lung (E and F).

Similar results were also obtained when dead cells from primary (noncultured) tissues were used instead of EL4. Intraperitoneal injection of necrotic lung homogenates elicited neutrophilic inflammatory responses, and injection of uricase reduced these responses significantly (Figure 6, C and D). Similarly, when necrotic lung from the intracellular uricase Tg animals was injected i.p. without exogenous uricase, we observed a similar reduction in inflammation relative to responses to WT lung (Figure 6, E and F). Therefore, uric acid depletion reduces inflammation to several types of dead cells and also in different anatomic locations (liver and peritoneum), although there are some quantitative differences in the degree of inhibition in these different situations.

Effect of uric acid depletion on inflammation due to sterile particulate or microbial stimuli. It was important to assess whether depleting uric acid was selectively affecting cell death–induced inflammation or whether it was generally immunosuppressive. To address this issue, we investigated to determine whether uric acid depletion would affect an inflammatory response in which uric acid should play no direct role. For this purpose, we injected sterile silica crystals, zymosan (yeast cell wall), or LPS i.p. into the various mouse models and quantified the ensuing inflammatory response. In untreated or buffer-treated WT animals, silica crystal, zymosan, and LPS elicit a neutrophilic inflammatory response similar to that of dead cells (Figure 7, A–C). There was no reduction in these inflammatory responses in the secreted or intracellular uricase Tg mice compared with WT animals (Figure 7, A–C). In fact, in the intracellular uricase Tg mice, the inflammatory responses to silica crystal, zymosan, and LPS were actually significantly increased for reasons that aren’t clear but are presumably somehow related to the hydrolysis of intracellular uric acid. In any case, this increased response to the sterile particulate or microbial stimuli makes the inhibition of responses to dead cells even more impressive. Zymosan-induced inflammation was also not decreased in mice treated with exogenous uricase or allopurinol compared with control animals (Figure 7, D and E). Collectively, these results are important because they indicate that uric acid depletion does not cause a generalized defect in generating an acute inflammatory response but instead is selectively affecting cell death–induced inflammation.

No decrease in neutrophil recruitment to silica crystals, zymosan, or LPS in uricase Tg mice or mice treated with rasburicase or allopurinol. Total number of neutrophils in the peritoneal cavity 15 hours after i.p. injection into WT or uricase Tg mice of 0.5 mg of silica crystal (A), 0.2 mg zymosan in uricase Tg mice (B), 100 ng of ultrapure LPS (C), or rasburicase-treated mice (D), or injection of zymosan into control, or allopurinol-treated mice (E). Means and SEM values are for combined data from 3 or more independent experiments. Number of mice used in (A): n =5 (PBS), n =11 (WT silica), n =8 (ssUOX Tg silica) 6 (intUOX Tg silica) (B): n = 4 (PBS) n = 15 (WT zymosan) n = 15 (ssUOX Tg zymosan) n = 17 (intUOX Tg zymosan) (C): n = 9 (PBS) n = 9 (WT LPS) n = 9 (ssUOX Tg LPS) n = 9 (intUOX Tg LPS) (D): n = 4 (PBS) n =15 (zymosan) n = 14 (zymosan + rasburicase) (E) n = 4 (PBS) n = 9 (zymosan) n = 9 (zymosan + allopurinol). *P < 0.05 NS, not significant versus WT silica (A), WT zymosan (B), or WT LPS (C).

The major finding in this report is that depletion of uric acid in vivo inhibits the inflammatory response to cell death. This inhibitory effect was seen in responses to necrotic liver, lung, and lymphoma cells and whether these responses were occurring in the tissues (liver) or the peritoneum. Therefore, uric acid is playing a role in inflammation to cell death for multiple different cell types and in different anatomical locations. Based on these findings, we predict that uric acid will play this role generally in multiple tissues.

Why does uric acid depletion inhibit inflammation? This is almost certainly due to the loss of uric acid rather than a pleiotropic effect of the method of its depletion because inhibition was seen regardless of whether uric acid levels were lowered by blocking its synthesis, lowering intracellular stores through enhanced catabolism, and/or eliminating it from the extracellular fluids, either acutely or chronically. In theory, uric acid could be influencing inflammatory responses in a few different ways. It is an antioxidant, so it is possible that eliminating it could affect inflammation through an alteration in redox potential ( 29 ). However, a generalized alteration in redox is unlikely to be a major mechanism of action because inflammation was inhibited whether extracellular uric acid levels were reduced (uricase treatment) or not (allopurinol treatment). Even more importantly, depletion of uric acid selectively inhibits inflammation to cell death rather than to other stimuli including microbial stimuli or irritant particles (silica). These latter findings are important because they argue that uric acid is not functioning as a general modulator of inflammation, as might occur if it were generally affecting redox potential. Instead, uric acid’s most likely mechanism of action is as a proinflammatory DAMP that is released from dying cells. This would be consistent with the known proinflammatory properties of monosodium urate (MSU). It would also explain the selective inhibition of cell death–induced inflammation by uric acid depletion. Moreover, it would explain why eliminating it intracellularly from dying cells or extracellularly after cell death would inhibit inflammation.

The model for uric acid acting as a proinflammatory DAMP is similar to the one that we have previously proposed ( 7 , 8 ). All cells contain intracellular pools of uric acid produced from the catabolism of purines, and these concentrations are quite high. Moreover, we found in the present study that after cell death there is a burst of uric acid production, as purines, presumably released by the action of nucleases on DNA and RNA, are converted by xanthine oxidase into uric acid. When this material is released into the extracellular fluids, the concentration of uric acid will exceed the saturation point in body fluids. Moreover, in the presence of the high levels of sodium in the extracellular fluids, the conditions should promote the nucleation of monosodium urate and, the hypothesis is, this is the biologically active form of the proinflammatory DAMP. While this model fits our data, it remains possible that uric acid is playing a role in some other way, e.g., by somehow controlling the release and/or activity of other DAMPs.

We had previously found that MSU and dead cells both stimulate sterile inflammation through the same final pathway ( 19 , 30 ). Both induce IL-1, and this mediator is essential for the neutrophilic inflammation in these responses. In contrast, IL-1 is not necessary for inflammation stimulated by microbial products ( 19 , 30 ). The present findings that dead cells work through uric acid may in part explain the common pathway.

Uric acid depletion inhibits cell death–induced inflammation only partially. Because uric acid is not completely eliminated in our studies, it is possible that its contribution to this response is being underestimated. To evaluate this point, it would be of interest to examine the effect of further reducing uric acid. We had hoped to achieve this with the uricase Tg mice, but despite analyzing multiple independent founders, we did not succeed in obtaining animals with more complete elimination. This raises the possibility that very low levels of uric acid levels are not permissive on a sustained basis. Despite being a “waste product,” there are several mechanisms that maintain high levels in vivo, including absorption from the diet and reabsorption through the kidney thus, uric acid might subserve some other useful function. In fact, we did find that while the secreted uricase Tg mice were healthy from birth through middle age, their survival past approximately 8 months was reduced the basis for this increased mortality will be studied and reported on in the future.

Given the large number of potential DAMPs, it is perhaps surprising that the elimination of a single one like uric acid had any effect at all. This could indicate that there are relatively few proinflammatory DAMPs and/or that uric acid is one of the major ones. It is also possible that uric acid plays a role that is different from other proinflammatory DAMPs uric acid is a small organic heterocyclic compound, while the other known DAMP candidates are proteins and therefore may work in different ways. If so, uric acid might provide additive or synergistic effects with other DAMPs. It would also not be surprising if different tissues have different contents of DAMPs, in which case a specific DAMP such as uric acid might play a greater or lesser role in different tissues. This might be one of the reasons in our experiments that uric acid depletion inhibits inflammation to different cell types to different degrees. It will be of interest to explore these possibilities in future studies.

The identification of proinflammatory DAMPs, such as uric acid, that play a role in vivo is of interest because these molecules are potential therapeutic targets. The inflammation stimulated by cell death is known to complicate acute ischemic and/or toxic injury to a number of organs, including the heart, liver, and lung, and is thought to also contribute to the pathogenesis of chronic diseases associated with cell death ( 5 , 31 , 32 ). Therefore, it could potentially be of therapeutic benefit to block this sterile inflammatory response. To this end, inhibiting the action of proinflammatory DAMPs would be attractive because it should selectively inhibit inflammation to cell death without affecting the ability of the host to mount inflammatory responses in other settings such as infection. Based on the present results, it will be of interest to test this concept with uric acid, especially because there are already uric acid–lowering drugs available that are well tolerated and approved for use in humans.

Chemicals and biologicals. Uric acid, allopurinol, acetaminophen, and zymosan were purchased from Sigma-Aldrich. UltraPure LPS was obtained from Invivogen (LPS-SM). Rasburicase (EC, recombinant uricase from Aspergillus flavus) was obtained from sanofi-aventis. Goat anti-uricase polyclonal antibody was from Santa Cruz Biotechnology Inc. Monoclonal anti–α-tubulin was from Abcam. Ly6G-FITC, anti-CD3e (145-2C11), and streptavidin-APC were from BD Bioscience. Biotinylated anti-7/4 was from AbD Serotec. Anti-CD45R (B220, clone RA3-6B2) and F4/80 (clone BM8) were from eBioscience.

Animals and cell lines. C57BL/6 and B6129 mice were purchased from Jackson Laboratory. To produce uricase Tg mice, a full-length cDNA for uricase was cloned from murine liver by RT–PCR. A secreted form of this enzyme was constructed by addition of signal sequence from adenovirus gp19K to the N terminus of the uricase cDNA. Secreted or unmodified (intracellular) uricase cDNAs were subcloned into pCAGGS ( 26 ) and injected into fertilized eggs from C57BL/6 mice that were then implanted into female mice to produce Tg mice. Primers used for genotyping were 5′-CAGCCATTGCCTTTTATGGT-3′ and 5′-CAGGACGTGCACAGTGTTCT-3′. The presence of the uricase transgene was further verified by Western blot of tissue lysates and measuring uricase activity in blood samples. All animal protocols were approved by the University of Massachusetts Medical School Animal Care and Use Committee. EL4 cells were maintained in RPMI with 10% FCS and antibiotics (Invitrogen) and tested negative for mycoplasma contamination (Lonza).

Uricase activity assay. To measure uricase activity in tissues, organs were harvested from mice and mixed with 4 times their weight in lysis buffer (20 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, and 0.1% SDS, complete protease inhibitor Roche Diagnostics GmbH) and homogenized with a tissue tearer followed by ultrasound sonication. After centrifugation at 15,000 g for 10 minutes, clarified supernatant was collected and assayed for protein concentration by the bicinchoninic acid method (Thermo Scientific). Lysate (20 μl) was added to 1 ml of uric acid solution (1.0 of OD292 in 0.1 M boric acid, pH 8.5) and incubated at 37°C. Reduction of OD292 of the uric acid solution was measured by spectrophotometer.

Western blot. Cleared lysates prepared as described above were measured for protein concentration, and a total of 10 μg of protein was fractionated by SDS–PAGE on a 10% gel, transferred to nitrocellulose membrane, and blotted with antibodies against uricase or α-tubulin.

Uric acid measurement. To measure uric acid in peritoneal cavity, the peritoneal cavity of mice was lavaged with 6 ml of physiological saline. Four ml of peritoneal lavage fluid was lyophilized and resuspended in 150 μl of water. The reconstituent was passed through an ultrafiltration membrane (3K cutoff Pall Life Sciences), and the filtrate was assayed for uric acid content (Bioassay Systems). The concentration of uric acid was also verified by HPLC using Superdex 200 (GE Healthcare). To measure uric acid concentration in tissues, the tissue lysates prepared as described above were kept cold or incubated at 37°C for the indicated time, passed through an ultrafiltration membrane, and assayed for uric acid concentration the concentrations reported in the figures are those in the lysates, which are much lower than those actually in cells ( 13 ) due to dilution in buffer.

Cell injury induction. EL4 cells were washed 5 times with PBS and resuspended in PBS at 10 million cells/50 μl. Cells were heat-shocked at 45°C for 10 minutes and incubated at 37°C for 5 hours. Mouse lung was washed 5 times with PBS the harvested lung was weighed and 5 times its weight in PBS was added, and then it was homogenated with a metal blade tissue tearer followed by 5 freeze-thaw cycles and 37°C incubation for 5 hours. Nine μg of rasburicase was added to 150 μl of cell suspension or lung homogenate at the start and end of a 5-hour incubation period.

Acetaminophen treatment and MPO assay. Mice were fasted for 18 hours and were then injected i.p. with 300 mg/kg acetaminophen (APAP) or 20 ml/kg PBS. Eighteen hours after APAP administration, blood was drawn for serum collection and ALT assay (Synchron LX Systems Beckman Coulter), and mice were euthanized to obtain liver tissues for MPO activity assay MPO is a neutrophil enzyme and a marker that is well validated for quantifying neutrophil infiltration into tissues ( 33 ). There were no difference in the MPO activities in the neutrophil in UOX Tg mice (Supplemental Figure 6B). In some experiments, mice were treated either with 18 μg of rasburicase i.p. at the same time as APAP treatment or with 1 week of allopurinol (10 mg/kg/d) i.p. injection daily before APAP treatment. After perfusion with PBS, 100 mg of liver was homogenized in 1 ml of MPO buffer (50 mM Na2HPO4, pH 5.4, 0.5% hexadecyl trimethyl ammonium bromide, 10 mM EDTA) using a tissue tearer and sonication. After centrifuge at 2500 g, 15 minutes at 4°C, 25 μl of supernatant was added with 25 μl of assay buffer (16.7 mg/ml of o-Dianisidine in 50 mM Na2HPO4, pH 5.4) and 200 μl of development solution (30 μl of 31.1% H2O2 per 10 ml of 50 mM Na2HPO4, pH 5.4). The OD450 change was measured using Bio-Rad 3550 microplate reader, and the rate between time 0 and 2 minutes 20 seconds was calculated.

Preparation of hepatic leukocytes and analysis by flow cytometry. After 18 hours of administration of APAP, livers were perfused with HBSS and liver digestion buffer (HBSS, 1.25 mM CaCl2, 4 mM MgCl2, 0.05% collagenase type IV, 0.028% DNAseI). Livers were homogenized and incubated in the digestion buffer for an hour at 37°C and passed through a 70-μm filter. Nonparenchymal cells were collected from the supernatant of centrifugation at 30 g and further purified from the interface in 22% of Opti Prep and RPMI 1640. The cells were stained with Ly6G and 7/4 antibodies to detect neutrophils and monocytes, anti-F4/80 antibody for macrophages, anti-CD3 antibody for T cells, and B220 antibody for B cells. The absolute number of cells was quantified using a flow cytometer with high-throughput sampler (BD Bioscience). Data were acquired by CellQuest software (BD Biosciences) and analyzed by FlowJo software (Tree Star Inc.).

Measurement of neutrophil recruitment to peritoneal cavity. To analyze the neutrophil recruitment to necrotic cells, mice were administered i.p. with 30 million cells of necrotic EL4 cells or 150 μl of lung homogenate. 0.2 mg of zymosan or 100 ng of pure LPS was also used to induce peritonitis. Fifteen hours after injection, animals were euthanized by CO2 exposure, and their peritoneal cavities were washed with 6 ml of RPMI with 10% FCS containing 3 mM EDTA and 10 U/ml heparin. The cells were stained with mAbs Ly-6G–FITC and 7/4-biotin for 30 minutes at 4°C in the presence of mAb 2.4G2 (FcγRIIB/III receptor blocker). The cells were further incubated with streptavidin-APC. Following staining, cells were washed with PBS with 2% FCS and resuspended with the original volume of the starting material. The total number of neutrophils or monocytes in the peritoneal exudate cells (PECs) was determined by the count of Ly-6G + 7/4 + or Ly-6G – 7/4 + cells in 100 μl of stained cells with a flow cytometer, respectively (FACSCalibur with high-throughput sampler). The results of the quantification of neutrophils in this assay correlates well with MPO levels in lysates of the lavage fluid (Supplemental Figure 6A).

Histology. 5-μm thickness sections from formalin-fixed and paraffin-embedded liver were cut on a microtome. To detect neutrophils, sections were stained with naphthol AS-D chloroacetate esterase according to manufacturer’s instructions (Sigma-Aldrich). Neutrophil number was counted in 10 high-power fields (×400) in each mouse using a Nikon Eclipse E800 Microscope. Ten low-power field (×100) pictures of each mouse were obtained from H&E-stained sections and the necrotic area quantified with ImageJ 1.42q (NIH) software.

Statistics. Statistical analysis in each independent experiment was performed with an unpaired, 2-tailed Student’s t test. Data are reported as mean ± SEM. One-way ANOVA and Dunnett’s multiple comparison post test were used to compare the means of multiple groups to the control group. P < 0.05 was considered statistically significant.

We thank Karen Dresser, Sharlene Hubbard, Matthew Janko, Zubin Patel, and Dipti Karmarkar for technical assistance. This work was supported by grants to K.L. Rock from the NIH and the American Asthma Foundation. Funding for this study was partially provided by sanofi-aventis US to H. Kono. Core resources supported by the Diabetes Endocrinology Research Center grant DK32520 were also used.

Conflict of interest: This research is partially funded from sanofi-aventis (to H. Kono).

Citation for this article: J Clin Invest. 2010120(6):1939–1949. doi:10.1172/JCI40124.

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The generation of point mutants using single-stranded DNA oligos and CRISPR/Cas9 genome editing reagents is an emerging technology in zebrafish and other animal model systems. The use of this technique enables single-nucleotide precision of genome editing experiments and will allow generation of specific disease models and precise mutational analysis of biological processes. Despite their obvious promise, point mutation knock-ins remain inefficient in zebrafish and the methods for testing their efficiency remain laborious or not easily accessible to many labs, such as sequencing individual plasmid clones by Sanger sequencing and NGS of PCR amplicons ( 11, 30). In an early TALEN-based knock-in study in zebrafish, restriction sites were introduced into specific genomic sites and shown to be digested by the corresponding enzyme ( 7, 8), but this approach has not yet been shown to work for CRISPR-based knock-ins in zebrafish. The restriction site introduction as a means of genotyping knock-ins is potentially attractive but these will have to be silent mutations in protein coding genes rather than insertions of complete sites. Moreover, PCR products with silent mutations may behave differently than those with added sequences. In the knock-in studies we describe here, we introduced missense mutations into tp53, cdh5 and lmna genes as well as synonymous mutations in PAM or sgRNA sites to prevent Cas9-mediated cutting or to introduce restriction sites for tp53 knock-ins. Restriction enzymes initially failed to genotype knock-in injected embryos, but were successful at genotyping of F1 heterozygous knock-in embryos. This discrepancy can be explained by the fact that in late PCR cycles, the strands of different PCR products can be randomly shuffled, from which follows that the fraction of PCR products having both strands containing knock-in mutations has a quadratic dependence on the knock-in allele frequency. Thus, at low (1 and 3%) knock-in rates (x), only a very small fraction (x 2 ) (0.01 and 0.09%) of total amplicon products will contain fully complementary strands and become digested. By contrast, in knock-in heterozygotes (50% allele frequency), 25% of PCR product can be digested, which was fully consistent with our results. We therefore switched to allele-specific PCR strategy to detect point mutations in all of our knock-ins and have shown that it is very sensitive to knock-in presence at allele frequencies <0.5%. Similar detection strategies were previously used for epitope-tagging knock-ins ( 9, 10), where one of the primers was specific to the tag inserts and the other was outside of the donor oligo region. Epitope tagging detection PCRs and point mutation AS-PCR assays are conceptually similar. However, the relatively small number of nucleotide differences between the WT and knock-in alleles can make it hard to avoid background amplification of a knock-in assay PCR in WT genomic DNA. We employed a touchdown PCR ( 31) protocol to make our AS-PCR strategies more specific, which was essential to the success of some AS-PCR assays and improved others. AS-PCR was also recently used in a mouse study of CRISPR/Cas9-based point mutation knock-in approaches ( 32) supporting the universal utility of this approach.

Overall, our studies of point mutation knock-ins revealed three main methods of improving efficiency. The first method was to reduce the distance between the mutation and Cas9 cut site ( 13). The first application of AS-PCR also indicated that this strategy may be useful in zebrafish since tp53 knock-ins where the distances were 10 and 13 nt were much less efficient than the cdh5 knock-in where the mutation was located exactly at the cut site. This experiment, although suggestive, could be improved by systematic variation of the mutation position relative to the cut site. The second approach, namely the usage of asymmetric oligos emerged as the key knock-in optimization in zebrafish. Knock-in AS-PCRs showed a very strong stimulation using this strategy in the case of tp53 knock-ins and NGS confirmed the significance of this result and allowed us to measure the extent of stimulation (ca. 3- and 10-fold for R143H and R217H knock-ins, respectively). Another explanation for why asymmetric oligos may function better comes from a well-established model proposing that the protruding single-stranded 3′ regions result from resection of DSBs ( 33). The team who explored this model performed knock-ins with multiple 97-nt oligos with different homology arms and they found that the most efficient oligos were 97 nt in length and were designed with shorter homology arms (30 nt) complementary to the resected single-stranded DNA ends produced after DSBs ( 12). These 30–67 asymmetric oligos introduced knock-ins equally well on either side of the DSB, thus supporting the resection model much more than the original model proposed by Richardson et al. ( 14). Future work will be necessary to establish if stimulation by asymmetric oligos on either strand can be equally effective, but our study provides evidence and an example of how this can be accomplished. However, the stimulation of knock-in efficiency by anti-sense asymmetric ssODNs may not be universal. For example, Moreno-Mateos et al. did not find any difference in knock-in efficiency between sense or anti-sense asymmetric oligos corresponding to the same stretch of genomic DNA when using SpCas9, but they measured that the anti-sense oligos were typically more efficient than the sense ones when DNA was cut by Cpf1 regardless of homology arm lengths ratios ( 34). Thus, both the genomic site and the nature of the CRISPR-related nuclease may play a role in determining the editing efficiency. In the third optimization, we tested PS modification of oligo ends while performing an lmna R471W knock-in. To uncouple potential effects of this modification from those of anti-sense asymmetric oligos and to avoid possible binding of oligos to ssDNA regions at off-target sgRNA sites, we chose sense asymmetric oligos of 90 nt with or without two PS bonds at either end of the oligo. Indeed, the PS-modified oligo was significantly more efficient and consistent at introducing knock-ins than the standard DNA oligo. Previously, the group that developed PS-mediated knock-in stimulation could only identify some imprecise knock-in events in zebrafish and did not test for knock-in improvement ( 15). We believe that all of these new optimization methods have utility and may even have multiplicative effects when deployed simultaneously. Therefore, at this stage of genome editing technology development, it is advisable to test several versions of oligos incorporating desired optimizations as well as a non-optimized control oligo in order to determine if the optimized versions behave in the expected way.

In the process of genotyping knock-in founders we developed a general workflow to identify true knock-in founders. The unexpected result that emerged from sequencing single F1 embryos was that there were many false-positive or trans knock-in founders (25–78% of total founder number). These could be screened out by sequencing or restriction digests, but they also revealed a weakness of AS-PCR strategies, which can falsely produce a positive signal likely due to hybridization of single DNA strands from abortive PCR products from target and trans loci. A possible mechanism of trans knock-in origin most likely has to do with off-target sgRNA sites, into which the oligo can ligate. Interestingly, the proportion of trans knock-in founders was much lower in all of our sense knock-in strategies than in all of anti-sense knock-ins. Anti-sense asymmetric oligos may have some complementarity to ssDNA regions generated at off-target sites, but this possibility needs further investigation.

In conclusion, we have provided and validated strategies to optimize and enhance point mutation knock-in efficiency in zebrafish. Proximity of the knock-in mutations to the Cas9 cut sites and anti-sense asymmetric oligos were identified as the most effective optimizations. PS modifications also enhanced knock-in efficiency and improved consistency among different embryos. These optimizations were enabled by AS-PCR assays and NGS. Restriction sites introduced by silent mutation as part of the knock-in process were also very useful for point mutant genotyping but only when they were present at high frequency (e.g. at 50%). We also identified the phenomenon of trans knock-ins, which can be filtered out using digestions of restriction sites introduced with silent mutations. We envision that this work will make point mutation knock-in generation a straightforward procedure accessible to all zebrafish researchers and other model systems researchers given the universal applicability of the methodology for point mutation introduction and optimization across a variety of animal models.

Watch the video: MITSUBISHI # 68214 (November 2021).