Skip to main content
Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

DNA Sequence Variants in the Five Prime Untranslated Region of the Cyclooxygenase-2 Gene Are Commonly Found in Healthy Dogs and Gray Wolves

  • Noa Safra ,

    Affiliation Department of Population Health and Reproduction, School of Veterinary Medicine, University of California Davis, Davis, California, United States of America

  • Louisa J. Hayward,

    Affiliation Animal Health Trust, Newmarket, United Kingdom

  • Miriam Aguilar,

    Affiliation Department of Population Health and Reproduction, School of Veterinary Medicine, University of California Davis, Davis, California, United States of America

  • Benjamin N. Sacks,

    Affiliation Veterinary Genetics Laboratory, School of Veterinary Medicine, University of California Davis, Davis, California United States of America

  • Jodi L. Westropp,

    Affiliation Department of Veterinary Medicine and Epidemiology, School of Veterinary Medicine, University of California Davis, Davis, California, United States of America

  • F. Charles Mohr,

    Affiliation Department of Pathology, Microbiology and Immunology, School of Veterinary Medicine, University of California Davis, Davis, California, United States of America

  • Cathryn S. Mellersh,

    Affiliation Animal Health Trust, Newmarket, United Kingdom

  • Danika L. Bannasch

    Affiliation Department of Population Health and Reproduction, School of Veterinary Medicine, University of California Davis, Davis, California, United States of America


10 Apr 2018: Safra N, Hayward LJ, Aguilar M, Sacks BN, Westropp JL, et al. (2018) Correction: DNA Sequence Variants in the Five Prime Untranslated Region of the Cyclooxygenase-2 Gene Are Commonly Found in Healthy Dogs and Gray Wolves. PLOS ONE 13(4): e0195924. View correction


The aim of this study was to investigate the frequency of regional DNA variants upstream to the translation initiation site of the canine Cyclooxygenase-2 (Cox-2) gene in healthy dogs. Cox-2 plays a role in various disease conditions such as acute and chronic inflammation, osteoarthritis and malignancy. A role for Cox-2 DNA variants in genetic predisposition to canine renal dysplasia has been proposed and dog breeders have been encouraged to select against these DNA variants. We sequenced 272–422 bases in 152 dogs unaffected by renal dysplasia and found 19 different haplotypes including 11 genetic variants which had not been described previously. We genotyped 7 gray wolves to ascertain the wildtype variant and found that the wolves we analyzed had predominantly the second most common DNA variant found in dogs. Our results demonstrate an elevated level of regional polymorphism that appears to be a feature of healthy domesticated dogs.


Cyclooxygenase (Cox) is the key enzyme in the biosynthesis of prostaglandins, which play a significant role in health and disease in the kidney, gastrointestinal tract, and in the skeletal and ocular systems [1]. There are two isozymes of Cox: a constitutively active form, Cox1 and an inducible form, Cox2. Cox-2 expression is regulated by stimulating conditions such as inflammation, injury or pain and it is upregulated during disease stages [2]. Cox-2 sequence variants that might modify the levels of protein expression would be expected to influence disease phenotype, and Cox-2 has been tested for associations with over 60 disorders with more than 1000 articles citing this gene in PubMed [3].

A role for Cox-2 DNA variants in canine renal dysplasia has been proposed since sequence variants were identified within the 5’ UTR region of canine Cox-2 in affected dogs [4]. Canine renal dysplasia (RD) is a progressive heritable nephropathy affecting many dog breeds [518]. Clinically, RD is characterized by clinical signs associated with chronic renal failure [17, 19, 20]. The age of onset of clinical signs varies between 3 weeks to ~3 years of age [16], and disease severity ranges between individuals [21], making this disorder challenging to diagnose [19].

The gold standard for diagnosis relies on renal biopsy and histopathological examination of a surgical wedge or a necropsy sample. It has been suggested that in order to make a definitive diagnosis, the sample must include at least 100 glomeruli, and the complete architecture of the renal cortex has to be evaluated [18, 20, 21]. Histologically, renal dysplasia can demonstrate a range of structural changes and includes findings inappropriate to the developmental stage of the dog examined [22]. There are 5 main morphologic characteristics of canine renal dysplasia, and the presence of at least one of them is required in order to obtain the diagnosis. They include asynchronous differentiation of nephrons (immature “fetal” glomeruli and tubules); persistent mesenchyme; persistent metanephric ducts; atypical tubular epithelium; and dysontogenic metaplasia [21].

Obtaining a diagnosis of renal dysplasia without a renal biopsy is difficult. While ultrasonography has been suggested as a screening tool, the changes noted ultrasonographically are not pathognomonic for renal dysplasia and can be found with other renal disorders [23]. Detecting mild or sub-clinical cases becomes even more difficult for clients and for breeders wanting to screen their dogs. Consequently, sub-clinical cases go undetected and are used in breeding programs, which may explain the high frequency of RD within certain breeds like the Lhasa Apso and the Shih-Tzu [4]. The high incidence of disease within certain breeds is suggestive of a familial disease, and both an incompletely penetrant autosomal dominant mode and a recessive mode of inheritance have been proposed [24]. Based on the presence of DNA variants in the 5’UTR of Cox-2 in RD affected dogs, a commercially available DNA test has been offered to dog breeders ( However, the causative nature of these DNA variants has come into question [25].

In order to ascertain the frequency of sequence variants at the 5’UTR of canine Cox-2, we evaluated sequences from dogs free of renal disease, and from gray wolves; the dog’s direct wild ancestor [26, 27]. Our findings suggest that regional variants are a common finding in dogs and do not present a genetic risk for RD.

Materials and Methods

DNA samples (n = 22) from dogs (Canis familiaris), obtained from patients of the Veterinary Medical Teaching Hospital at UC Davis that underwent complete necropsy and did not have histological evidence of RD were analyzed. Additional samples (n = 46) came from dogs 6 years old or older that had normal blood urea nitrogen (BUN) and creatinine concentrations (values were recorded during routine serum biochemical analysis). Samples of Flat-Coated Retrievers (FCRs) (n = 84) from the DNA collection of the Kennel Club Genetics Centre at the Animal Health Trust (AHT) in Newmarket, UK (submitted to the AHT with owner consent for use in variable research projects), were also used. The cohort of 84 FCRs ranged in age from 5 months to 14 years at the time of sample submission, and each dog had unique parentage (168 different dams and sires are represented). These dogs were free of renal disease at the time of sample submission, according to owners. Additional samples (n = 12) came from gray wolves (Canis lupus) collected from the Yukon Territory, Canada. These samples were archived for over ten years and used previously [28]; no samples were collected specifically for the purpose of this study and no animals were killed specifically for this study. The wolf samples were genotyped to ensure that our analysis was restricted to independent (i.e. unrelated) wolves. A panel of 24 short tandem repeat (STR) markers representing 20 canine autosomes was used by the Veterinary Genetics Laboratory (VGL) as was previously described [29]. The gray wolf relatedness was estimated using a maximum likelihood approach [30], and individuals with estimated relatedness <0.15 were considered to be unrelated which left 7 wolves for the analysis. Genomic DNA was extracted from EDTA whole blood or tissue samples using Gentra Puregene DNA purification extraction kit (Qiagen, Valencia, CA) or the Nucleon Genomic DNA Extraction Kit (Tepnel Life Sciences, Manchester, UK). Buccal swab DNA was extracted using the QIAamp DNA Blood Midi Kit (Qiagen, Valencia, CA).

The published primers [4]: 1-F: 5'-TTGTCAAACAACTTGCAGCGAGCG; 1-R: 5'-ATCACCCAGCCGAGGAGTC that amplify 272 bases were used to genotype the FCRs (n = 84), and 2-F: 5′ GCCCTTTACCAGGTCAAACA; 2-R: 5′ GCGAGGACCCAACAGCTTAC that amplify 422 bases were used to genotype the rest of the samples (n = 68). Reaction mixes using primers 1F and 1R consisted of 0.2 mM dNTPs, 1x PCR buffer, 1x Q solution (Qiagen), 0.8 μM of each primer, 0.5 U HotStarTaq Plus DNA Polymerase (Qiagen), 2 μl genomic DNA, and ultrapure water to give a final volume of 12 μl. Cycling conditions for PCR were 3 minutes at 95°C, followed by 35 cycles of 30 seconds at 95°C, 30 seconds at 60°C, and 1 minute at 72°C, with a final extension of 10 minutes at 72°C. Products were sequenced using BigDye Terminator v3.1 on an ABI 3130xl Genetic Analyzer (Applied Biosystems). Sequence traces were visually analyzed using the Staden Gap4 package [31]. Primers 2F and 2R (Tm = 68°C) were used to generate PCR products using AccuPrime GC-Rich DNA Polymerase (AccuPrime high GC DNA polymerase kit; life technologies, Grand Island, NY 14072, USA). PCR cycle conditions, Sanger sequencing protocol and sequence analysis were previously described [32]. For heterozygous individuals, PCR products were cloned, sequenced, and analyzed as was previously described [32].


We investigated the presence of DNA sequence variants within the 5’UTR of the canine Cox-2 gene in FCRs reported free of renal disease by owners (n = 84) and in samples from dogs free of RD based on their clinical biochemistry or necropsy results (n = 68). Forty six dogs with serum BUN (reference range 6–31 mg/dL), creatinine (reference range 0.5–1.6 mg/dL) concentrations within the reference ranges, and no clinical signs associated with renal disease; and 22 dogs had necropsy results confirming absence of RD. The entire cohort consisted of 142 pure-bred dogs from 30 different breeds, and of 10 mixed-bred dogs (S1 Table).

The first coding ATG, according to dog mRNA sequence (GenBank: AY044905) represents position zero. The sequence variants identified upstream to position zero are described based on standard mutation nomenclature recommended by the Human Genome Variation Society (HGVS) [33], and in relation to CanFam3.1 sequence assembly. However, because of the repetitive high-GC content of the 5’UTR sequence of the Cox-2 gene, the exact base positioning of certain sequence variants could not be determined (Fig 1, S1 Table).

Fig 1. DNA sequence variants identified within the 5’UTR of the Cox-2 gene in 152 dogs.

Base positions upstream of the ATG translation initiation site (position zero) are noted on the left. *The exact base position for the duplicated sequence could not be determined. The sequence variant most commonly found in gray wolves is boxed.

Table 1 lists the variants found in the cohort of 68 dogs by nucleotide position, variant type and haplotype frequency. S2 and S3 Tables list the haplotypes observed within the subgroups of dogs without evidence of azotemia (n = 46), and dogs that were found free of RD by necropsy (n = 22). There were no statistically significant differences between the frequencies of the variants found within these two groups (Fisher exact test p value = 0.658842). The majority of the dogs (n = 44) had at least one haplotype identical to the CanFam3.1 assembly sequence, and the frequency of the CanFam3.1 haplotype was 0.4 in this sample set. Sequence variant Cox2–77_-76ins12 was the second most common at a frequency of 0.3. Recombined haplotypes and novel variants were commonly observed and a total of 17 haplotypes were observed in the 68 dogs. Listed in Table 1 are 11 genetic variants which had not previously been described.

Table 1. 5’UTR Cox-2 variants found in 68 dogs free of renal disease*.

Breeds that were reported to have had 5’UTR Cox-2 variants concurrent with RD [4], were represented in this cohort by the Boxer (n = 4), with variants Cox2–72_-67del6, Cox2 -42T>C and Cox2–37_-27del11; Bernese Mountain dog (n = 2) Cox2–77_-76ins12, Cox2–72_-67del6, Cox2 -42T>C and Cox2–37_-27del11; Australian Shepherd (n = 1) Cox2–77_-76ins12; Labrador Retriever (n = 2) Cox2–77_-76ins30 and Cox2–72_-67del6, Cox2 -42T>C and Cox2–37_-27del11; Yorkshire Terrier (n = 1) Cox2–77_-76ins30; Rottweiler (n = 1) Cox2–77_-76ins24; and German Shepherd dogs (n = 10) Cox2–72_-67del6, Cox2 -42T>C and Cox2–37_-27del11(n = 3). We sequenced unaffected Shetland Sheepdog (n = 9) and found that all of the individuals tested had regional variants, and dogs from this breed had haplotypes that recombine variants Cox2–77_-76ins30/24/12 and Cox2–72_-67del6, Cox2 -42T>C and Cox2–37_-27del11 (S1 Table).

The FCR breed was represented by the largest cohort (n = 84). The sequence variants identified within FCRs are listed in Table 2 by nucleotide position and haplotype frequency. The most frequent haplotype documented within the FCRs was CanFam3.1 assembly sequence at allele frequency of 0.50, followed by a novel haplotype found in FCRs, Cox2-72_-67del6; -42T>C; -27T>C, at a frequency of 0.38. Less frequent were a second novel haplotype unique to FCRs, Cox2-77_-76ins24; -67_-66dup6 (0.06), and Cox2-72_-67del6; -42 T>C; -37_-27del11 (0.06). The novel haplotypes found in FCRs brought the total number of haplotypes observed in dogs to 19.

Table 2. 5’UTR Cox-2 variants found in 84 Flat-Coated Retrievers.

In order to clarify the ancestral state of the 5’ UTR sequence of canine Cox-2, 12 DNA samples from gray wolves were sequenced. Based on STRs, a matrix of maximum likelihood estimates of relatedness revealed that 7 of the 12 wolves were unrelated. The 5’UTR haplotypes of the 7 unrelated wolves are presented in Table 3. The most common haplotype observed within wolves was sequence variant Cox2–77_-76ins12 at allele frequency of 0.72. A single wolf was heterozygous for a novel variant Cox2–77_-76ins19, and two novel SNPs were identified in wolves; Cox2 -67G>T, and Cox2 -62T>G (Table 3). A single wolf was heterozygous for CanFam3.1 assembly sequence. The five remaining wolves shared ancestry and all of them were homozygous for Cox2–77_-76ins12. The wolves sampled came from a genetically diverse population based on high heterozygosity values observed for the STR markers (S4 Table).

Table 3. 5’UTR Cox-2 variants found in 7 Alaskan gray wolves.

We next compared Cox-2 5’UTR sequences across species. mRNA sequences from Human (Homo sapiens; accession: U04636), Horse (Equus caballus; AF027334), Cow (Bos taurus; DT845401), Mouse (Mus musculus; BB653429), and Chicken (Gallus gallus; NM_001167719) were aligned together with genomic sequences from Wolf (Canis lupus; KP296709), and CanFam3.1 sequence (Canis lupus familiaris; canFam3_dna range = chr7:19674506–19674874). The alignment, shown in Fig 2, demonstrates that a conserved regional sequence includes ~12 additional nucleotides when compared to the CanFam3.1 assembly.

Fig 2. Alignment of DNA sequences upstream of the ATG of the Cox-2 gene from Human, Horse, Cow, Wolf, CanFam3.1 assembly, Mouse and Chicken.

The presumed translation initiation ATG is boxed. A DNA variant (deletion of 12 nucleotides) is observed in CanFam3.1 sequence when compared to other organisms (double headed arrow).


The genomic region immediately upstream of the Cox-2 gene was sequenced in 3 groups of dogs free of clinical signs of renal disease as well as in gray wolves. The first group of dogs consisted of unaffected individuals cleared by necropsy. To increase the sample size we included non-azotemic dogs older than 6 years of age because RD is usually documented in juvenile animals [16] [23]. However, there are reports of older cases [8], and age of onset could be a breed specific trait. The third group of dogs consisted of FCRs free of clinical signs according to owner questionnaire without medical records. While a limitation of the study is the incomplete phenotyping of some of the dogs, the results were similar between the groups.

Approximately half of the haplotypes observed in dogs represented CanFam3.1 assembly, while the other half featured a high degree of polymorphism including insertions, duplications, deletions and SNPs. Novel variants were a common observation and a total of 11 new variants were described. Surprisingly, the wolves sampled showed lower levels of regional polymorphism when compared to dogs and 11 of the 12 wolves had sequence variant Cox2–77_-76ins12. While the number of wolf samples available for this study is insufficient to draw final conclusions, the presence of fewer wolf haplotypes than dogs’ was an unexpected result. This is because the wolves showed high levels of genetic diversity at 24 genome-wide STR markers and because the wolf genome is more polymorphic than the dog’s [26]. Additional sequences from wolves may elucidate this observation.

Across species alignment of the regional DNA sequence revealed that a haplotype with additional 12 bases, comparable to -77_-76ins12, is conserved between gray wolves, Human, Horse, Cow, Mouse and Chicken. This suggests that this is the wildtype haplotype, particularly since this was also the second most common haplotype found in non-azotemic dogs. The wild-type sequence is not expected to increase the likelihood of RD, challenging the claim that this variant and/or other regional sequence variants harbor susceptibility to RD in dogs. Based on our results, we propose that additional studies are needed in order to uncover RD susceptibility loci to assist dog breeders in selecting against RD in predisposed breeds.

Currently, the functional significance of the regional sequence variants is unknown. The canine Cox-2 promoter has been identified and employed successfully by Rachakonda et al (2008) and Rai et al (2011) in in-vitro arthritis models using canine chondrocytes [34, 35]. Only a short region of ~64 bases at the 3' end of this canine Cox-2 promoter overlaps with the region containing the variants reported by Whiteley et al (2011) [4]. Although the overlapping region contains sequences that may correspond to SP1 sites (CpG islands), Nunez et al (2011) have identified the main regulatory regions to be further upstream in the human Cox-2 gene [36]. Cox-2 basal level expression is variable among individual dogs [37], but variability in expression is also noted within a species with fewer documented 5’UTR sequence polymorphism such as people [38, 39].

An explanation for the presence of multiple DNA variants immediately upstream to the presumed translation initiation site of canine Cox-2 in dogs requires further investigation. Sequence polymorphisms within the 5’UTR of other canine genes have been studied and show moderate variation comparing to the various haplotypes observed within the canine Cox-2 gene. Young and Bannasch (2008) acquired 5’UTR sequences from 24 unrelated dogs and revealed a single SNP each in the SHOX and RMRP genes [40], and Shirazi-Beechey et al (2013) screened 60 dogs to reveal 3 SNPs within the 5’UTR region of the SGLT1 gene [41]. It is possible that this genomic region represents a recombination hotspot, as Auton et al (2013) studied recombination hotspots in dogs and observed elevated rates of recombination around transcription start sites. It has been reported that CpG content is strongly associated with canine recombination and that the most recombinogenic repeats are low-complexity with high levels of GC and CpG content [42], as found in this region.

Ethic Statement

Samples donated to the AHT with owner consent for use in variable research projects from Flatcoated Retrievers were submitted in the form of buccal swabs or blood. Buccal swabs were collected by owners using a non-invasive method. Blood samples were the remnants of blood drawn by veterinarians for routine and/or diagnostic veterinary purposes, and were not collected specifically for the purposes of research.

For the portion of the study carried out in the University of California, the sample collection protocol was approved by the University of California, Davis Animal Care and Use Committee (protocol #16892), with owner consent. Additional samples came from blood samples that were drawn for medical purposes at the Veterinary Teaching Hospital, UC Davis. DNA was extracted from unused portions of the samples. Tissue samples from dogs that underwent complete necropsy were used for research purposes with owner’s consent.

Gray wolves (Canis lupus) samples collected from the Yukon Territory, Canada, were archived for over 10 years and used previously [28]. No samples were collected specifically for the purpose of this study and no animals were killed specifically for this study.

Supporting Information

S1 Table. Breed representation of the 19 haplotypes found within the 5’UTR of Cox-2 in 152 dogs.

*novel variants.


S2 Table. 5’UTR Cox-2 variants found in 46 dogs that had normal blood urea nitrogen and creatinine levels and no history of renal disease.

*novel variants.


S3 Table. 5’UTR Cox-2 variants found in 22 dogs that underwent complete necropsy and were clear of RD.

*novel variants.


S4 Table. Heterozygosity values observed in 12 Alaskan gray wolves using the canine parental verification panel at the *VGL.

PIC: Polymorphism Information Content (a value derived to measure the infomativeness of a genetic marker). *


Author Contributions

Conceived and designed the experiments: DLB JLW CSM BNS NS. Performed the experiments: LJH NS MA BNS. Analyzed the data: MA NS DLB LJH BNS CSM FCM. Contributed reagents/materials/analysis tools: JLW FCM. Wrote the paper: NS LJH CSM DLB.


  1. 1. Doré M. Cyclooxygenase-2 Expression in Animal Cancers. Veterinary Pathology Online, 2011. 48(1): p. 254–265.
  2. 2. Gierse JK, Hauser SD, Creely DP, Koboldt C, Rangwala SH, Isakson PC, et al. Expression and selective inhibition of the constitutive and inducible forms of human cyclo-oxygenase. Biochem J, 1995. 305 (Pt 2): p. 479–84. pmid:7832763
  3. 3. Thierry-Mieg D, Thierry-Mieg J. AceView: a comprehensive cDNA-supported gene and transcripts annotation. Genome Biol, 2006. 7 Suppl 1: p. S12 1–14. pmid:16925834
  4. 4. Whiteley MH, Bell JS, Rothman DA. Novel allelic variants in the canine cyclooxgenase-2 (Cox-2) promoter are associated with renal dysplasia in dogs. PLoS One, 2011. 6(2): p. e16684. pmid:21346820
  5. 5. de Morais HS, DiBartola SP, Chew DJ. Juvenile renal disease in golden retrievers: 12 cases (1984–1994). J Am Vet Med Assoc, 1996. 209(4): p. 792–7. pmid:8756882
  6. 6. Abraham LA, Beck C, Slocombe RF. Renal dysplasia and urinary tract infection in a Bull Mastiff puppy. Aust Vet J, 2003. 81(6): p. 336–9. pmid:15080453
  7. 7. Felkai C, Voros K, Vrabely T, Vetesi F, Karsai F, Papp L. Ultrasonographic findings of renal dysplasia in cocker spaniels: eight cases. Acta Vet Hung, 1997. 45(4): p. 397–408. pmid:9557317
  8. 8. Hoppe A, Karlstam E. Renal dysplasia in boxers and Finnish harriers. J Small Anim Pract, 2000. 41(9): p. 422–6. pmid:11023130
  9. 9. Lobetti RG, Pearson J, Jimenez M. Renal dysplasia in a Rhodesian ridgeback dog. J Small Anim Pract, 1996. 37(11): p. 552–5. pmid:8934429
  10. 10. Ohara K, Kobayashi Y, Tsuchiya N, Furuoka H, Matsui T. Renal dysplasia in a Shih Tzu dog in Japan. J Vet Med Sci, 2001. 63(10): p. 1127–30. pmid:11714030
  11. 11. Olenick CL. Congenital renal dysplasia and psychogenic polydipsia in a Bernese mountain dog. Can Vet J, 1999. 40(6): p. 425–6. pmid:10367161
  12. 12. Schulze C, Meyer HP, Blok AL, Schipper K, van den Ingh TS. Renal dysplasia in three young adult Dutch kooiker dogs. Vet Q, 1998. 20(4): p. 146–8. pmid:9810631
  13. 13. Roels S, Schoofs S, Ducatelle R. Juvenile nephropathy in a Weimaraner dog. J Small Anim Pract, 1997. 38(3): p. 115–8. pmid:9097243
  14. 14. Booth K. A case of juvenile nephropathy in a Newfoundland dog. Vet Rec, 1990. 127(24): p. 596–7. pmid:2075690
  15. 15. DiBartola SP, Chew DJ, Boyce JT. Juvenile renal disease in related Standard Poodles. J Am Vet Med Assoc, 1983. 183(6): p. 693–6. pmid:6629980
  16. 16. O'Brien TD, Osborne CA, Yano BL, Barnes DM. Clinicopathologic manifestations of progressive renal disease in Lhasa Apso and Shih Tzu dogs. J Am Vet Med Assoc, 1982. 180(6): p. 658–64. pmid:7068507
  17. 17. Morton LD, Sanecki RK, Gordon DE, Sopiarz RL, Bell JS, Sakas PS. Juvenile renal disease in miniature schnauzer dogs. Vet Pathol, 1990. 27(6): p. 455–8. pmid:2278134
  18. 18. Bruder MC, Shoieb AM, Shirai N, Boucher GG, Brodie TA. Renal dysplasia in Beagle dogs: four cases. Toxicol Pathol, 2010. 38(7): p. 1051–7. pmid:20884818
  19. 19. Lees GE. Congenital renal diseases. Vet Clin North Am Small Anim Pract, 1996. 26(6): p. 1379–99. pmid:8911024
  20. 20. Greco DS. Congenital and inherited renal disease of small animals. Vet Clin North Am Small Anim Pract, 2001. 31(2): p. 393–9, viii. pmid:11265498
  21. 21. Picut CA, Lewis RM. Microscopic features of canine renal dysplasia. Vet Pathol, 1987. 24(2): p. 156–63. pmid:3576910
  22. 22. Donald M, McGavin JFZ. Pathologic Basis of Veterinary Disease. 4 ed. Vol. 4th ed. 2006: Mosby. 1,488.
  23. 23. Seiler GS, Rhodes J, Cianciolo R, Casal ML. Ultrasonographic findings in Cairn Terriers with preclinical renal dysplasia. Vet Radiol Ultrasound, 2010. 51(4): p. 453–7. pmid:20806879
  24. 24. Bovee KC. “Renal Dysplasia in Shih Tzu Dogs”. Proceedings of the 28th World Congress of the World Small Animal Veterinary Association. 2003.
  25. 25. Expression of concern: Novel allelic variants in the canine cyclooxgenase-2 (Cox-2) promoter are associated with renal dysplasia in dogs. PLoS One, 2012. 7(11): p. e49703. pmid:23145184
  26. 26. Lindblad-Toh K, Wade CM, Mikkelsen TS, Karlsson EK, Jaffe DB, Kamal M, et al. Genome sequence, comparative analysis and haplotype structure of the domestic dog. Nature, 2005. 438(7069): p. 803–19. pmid:16341006
  27. 27. Larson G, Karlsson EK, Perri A, Webster MT, Ho SY, Peters W, et al. Rethinking dog domestication by integrating genetics, archeology, and biogeography. Proc Natl Acad Sci U S A, 2012. 109(23): p. 8878–83. pmid:22615366
  28. 28. Brown SK, Pedersen NC, Jafarishorijeh S, Bannasch DL, Ahrens KD, Wu JT, et al. Phylogenetic Distinctiveness of Middle Eastern and Southeast Asian Village Dog Y Chromosomes Illuminates Dog Origins. PLoS ONE, 2011. 6(12): p. e28496. pmid:22194840
  29. 29. Pedersen NC, Liu H, Theilen G, Sacks B. The effects of dog breed development on genetic diversity and the relative influences of performance and conformation breeding. Journal of Animal Breeding and Genetics, 2013. 130(3): p. 236–248. pmid:23679949
  30. 30. Kalinowski ST, Wagner AP, Taper ML. Relate: a computer program for maximum likelihood estimation of relatedness and relationship. Molecular Ecology Notes 2006. 6: p. 576–579.
  31. 31. Bonfield JK, Smith KF, Staden R. A new DNA sequence assembly program. Nucleic Acids Research, 1995. 23(24): p. 4992–4999. pmid:8559656
  32. 32. Safra N, Bassuk AG, Ferguson PJ, Aguilar M, Coulson RL, Thomas N, et al. Genome-Wide Association Mapping in Dogs Enables Identification of the Homeobox Gene, NKX2-8, as a Genetic Component of Neural Tube Defects in Humans. PLoS Genet, 2013. 9(7): p. e1003646. pmid:23874236
  33. 33. Ogino S, Gulley ML, den Dunnen JT, Wilson RB. Standard Mutation Nomenclature in Molecular Diagnostics: Practical and Educational Challenges. The Journal of Molecular Diagnostics, 2007. 9(1): p. 1–6. pmid:17251329
  34. 34. Rachakonda PS, Rai MF, Schmidt MFG. Application of inflammation-responsive promoter for an in vitro arthritis model. Arthritis & Rheumatism, 2008. 58(7): p. 2088–2097.
  35. 35. Rai MF, Graeve T, Twardziok S, Schmidt MF. Evidence for Regulated Interleukin-4 Expression in Chondrocyte-Scaffolds under In Vitro Inflammatory Conditions. PLoS ONE, 2011. 6(10): p. e25749. pmid:21991344
  36. 36. Nuñez F, Bravo S, Cruzat F, Montecino M, De Ferrari GV. Wnt/β-Catenin Signaling Enhances Cyclooxygenase-2 (COX2) Transcriptional Activity in Gastric Cancer Cells. PLoS ONE, 2011. 6(4): p. e18562. pmid:21494638
  37. 37. Dudley A, Thomason J, Fritz S, Grady J, Stokes J, Wills R, et al. Cyclooxygenase Expression and Platelet Function in Healthy Dogs Receiving Low Dose Aspirin. Journal of veterinary internal medicine / American College of Veterinary Internal Medicine, 2013. 27(1): p. 141–149. pmid:23278865
  38. 38. Hamajima N, Takezaki T, Matsuo K, Saito T, Inoue M, Hirai T, et al. Genotype Frequencies of Cyclooxygenease 2 (COX2) Rare Polymorphisms for Japanese with and without Colorectal Cancer. Asian Pac J Cancer Prev, 2001. 2(1): p. 57–62. pmid:12718655
  39. 39. Frelinger AL 3rd, Furman MI, Linden MD, Li Y, Fox ML, Barnard MR, et al. Residual arachidonic acid-induced platelet activation via an adenosine diphosphate-dependent but cyclooxygenase-1- and cyclooxygenase-2-independent pathway: a 700-patient study of aspirin resistance. Circulation, 2006. 113(25): p. 2888–96. pmid:16785341
  40. 40. Young AE, Bannasch DL. SNPS in the Promoter Regions of the Canine RMRP and SHOX Genes are not Associated with Canine Chondrodysplasia. Animal Biotechnology, 2008. 19(1): p. 1–5. pmid:18228171
  41. 41. Batchelor DJ, German AJ, Shirazi-Beechey SP. Relevance of sodium/glucose cotransporter-1 (SGLT1) to diabetes mellitus and obesity in dogs. Domestic Animal Endocrinology, 2013. 44(3): p. 139–144. pmid:23375266
  42. 42. Auton A, Ying RL, Jeffrey K, Oliveira K, Nadel J, Holloway JK, et al. Genetic Recombination Is Targeted towards Gene Promoter Regions in Dogs. PLoS Genetics, 2013. 9(12): p. e1003984. pmid:24348265