1

Tools for detection of Mycoplasma amphoriforme; a primary respiratory pathogen? 1

2

3

Clare L. Linga*, Katarina Oravcovab, Thomas F. Beattieb,c, Dean D. Creerd, Paul Dilworthe, Naomi 4

L. Fultonf, Alison Hardief, Michelle Munro,f Marcus Pond,g Kate Templeton,f David Webster,h 5

Sarita Workman,h Timothy D. McHugh,a Stephen H. Gillespie,b# 6

7

Centre for Clinical Microbiology, Department of Infection, Royal Free Campus, University College 8

London, London, UKa; School of Medicine, University of St. Andrews, St Andrews, UKb; Royal 9

Hospital for Sick Children, Edinburgh, UKc; Respiratory Medicine, Barnet General Hospital, 10

London, UKd; University College London Medical School, University College London, London, 11

UKe; Department of Medical Microbiology, Royal Infirmary, Edinburgh, UKf; Centre for Infection 12

and Immunity, St George’s University of London, London, UKg; Department of Immunology, Royal 13

Free London NHS Foundation Trust, London, UKh 14

15

Running Title: Mycoplasma amphoriforme detection 16

17

#Address correspondence to Stephen H. Gillespie, shg3@st-andrews.ac.uk 18

*Present address: Clare L. Ling, Shoklo Malaria Research Unit, Mahidol-Oxford Tropical Medicine 19

Research Unit, Faculty of Tropical Medicine, Mahidol University, Mae Sot, Thailand 20

21

22

23

JCM Accepts, published online ahead of print on 29 January 2014J. Clin. Microbiol. doi:10.1128/JCM.03049-13Copyright © 2014, American Society for Microbiology. All Rights Reserved.

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

2

ABSTRACT 24

Mycoplasma amphoriforme is a recently described organism isolated from the respiratory tract of 25

patients with immunodeficiency and evidence of chronic infection. Novel assays for the molecular 26

detection of the organism by real-time quantitative PCRs (qPCR) targeting the uracil DNA 27

glycosylase (udg) gene and 23S ribosomal DNA are described. The analytical sensitivities are 28

similar to the existing conventional 16S rDNA M. amphoriforme PCR with the advantage of being 29

species-specific, rapid and quantitative. Using these techniques we demonstrate the presence of this 30

organism in 17 (19.3%) primary antibody deficient (PAD) patients, 4 (5%) adults with lower 31

respiratory tract infection, 1 (2.6%) sputum sample from patients attending a chest clinic, 23 32

(0.21%) samples submitted for viral diagnosis of respiratory infection, but not in normal adult 33

control subjects. These data show the presence of this microorganism in respiratory patients and 34

suggest that M. amphoriforme may infect both immunocompetent and immunocompromised 35

subjects. Further studies to characterise this organism are required and this report provides the tools 36

that may be used by other research groups to investigate its pathogenic potential. 37

38

39

INTRODUCTION 40

Mycoplasma amphoriforme was first isolated in 1999 from a patient with primary antibody 41

deficiency (PAD) with chronic bronchitis. It has also been isolated subsequently from both 42

immunocompromised and immunocompetent patients with respiratory tract infections (RTI) in 43

London, Denmark, France and Tunisia (1-3). Based on 16S rRNA sequencing M. amphoriforme 44

belongs to the same phylogenetic group as other human pathogenic Mycoplasma species, the 45

pneumoniae group (1, 2). The closest species phylogenetically for which there is a whole genome 46

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

3

sequence is Mycoplasma gallisepticum, a bird pathogen. Phenotypic studies have demonstrated that 47

M. amphoriforme has features in common with this group including: gliding motility, a protruding 48

polar tip resembling that of M. gallisepticum and a cytoskeletal structure at its polar tip with 49

homology to that of M. pneumoniae's attachment organelle (1, 4). 50

51

To understand the role that this novel agent plays in human health, better laboratory tools are 52

required. M. amphoriforme is fastidious, requiring specialised media for cultivation and it takes 53

approximately two weeks for colonies to appear on agar. The colonial morphology resembles 54

granular droplets making the detection difficult as they can blend into the sample matrix and be 55

overlooked. This paper reports the development and evaluation of two real-time PCR (qPCR) 56

assays, an assay targeting M. amphoriforme’s uracil DNA glycosylase (udg) gene and an assay 57

targeting the variable region of 23S rDNA (23S) that is unique to M. amphoriforme. The new qPCR 58

assays were compared with a previously reported 16S rDNA assay (16S) (2) and used to test a range 59

of human samples from the UK. 60

61

62

63

MATERIALS AND METHODS 64

65

Patients, samples and ethical approval 66

Clinical samples from two hospitals were used in this study, the Royal Free London NHS 67

Foundation Trust (RFL), Hampstead and the Royal Infirmary of Edinburgh (RIE). The approvals 68

were obtained from the Ethics committee of the RFL Hampstead and from the Lothian Regional 69

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

4

Ethics Committee (08/S11/02/2) to retain information during anonymisation for epidemiological 70

purposes. 71

72

From 19th October 2000 to 6th September 2005 sputum samples were collected from PAD patients 73

attending the dedicated Primary Immunodeficiency Clinic at the RFL. They attended for either a 74

routine appointment or in cases of clinical deterioration. A sputum sample was collected from any 75

patient with a productive cough and sent for microbiological investigation, including Mycoplasma 76

amphoriforme detection. The age range of all the PID patients tested was 18 to 79 with an average 77

age of 44 years. 78

Sputum and/or throat swab samples were collected from adult patients attending the RFL Chest 79

Clinic and from adult patients (≥18 years) with lower respiratory tract infections (LRTI) that were 80

recruited from two general practices with a multi-ethnic patient population of 15 000 from social 81

classes I–V as described previously (5). All LRTI patients were surgery attendees; no recruitment 82

was undertaken out of hours or on home visits. Acute LRTI was defined as a new or worsening 83

cough and at least one other lower respiratory tract symptom for which there was no other 84

explanation, present for 21 days or less (6, 7). Patients were excluded if they had underlying chronic 85

suppurative lung disease (defined as bronchiectasis, lung abscess or empyema), tuberculosis, 86

immunodeficiency, or previous study participation (three weeks). Age, sex, and season matched 87

controls were recruited from general practice patients attending for non-respiratory and non-88

infective illnesses as well as other healthy volunteers with no history of respiratory tract symptoms 89

for two months prior to recruitment using the same exclusion criteria as previously described for 90

patients (5). 91

Anonymised respiratory samples including sputum, nasopharyngeal secretions and throat swabs 92

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

5

collected at hospital and primary care settings in South East Scotland referred to the RIE Specialist 93

Virology Centre (SVC) from adults and children with suspected respiratory infection and submitted 94

for viral diagnosis were tested. The age of the patients ranged from 0-96 years with a mean of 19.91 95

years. The stored data for these samples included age band, partial postcode, any recorded 96

symptoms or clinical information, referral source, month of sample collection, and results of other 97

virological testing of the sample. 98

99 Control organisms 100

The following control organisms were used to test the specificity of the assays: M. amphoriforme 101

NCTC 11740 and Mycoplasma pneumoniae ATCC 5167 (Mycoplasma Experience Ltd. UK); 102

Mycoplasma testudinis NCTC 11701 and Mycoplasma alvi ATCC 29626 (Leahurst, UK); 103

Acholeplasma laidlawii ATCC 23206, Mycoplasma buccale ATCC 23636, Mycoplasma faucium 104

ATCC 25293, Mycoplasma fermentans ATCC 19989, Mycoplasma genitalium ATCC 33530, 105

Mycoplasma hominis ATCC 23114, Mycoplasma orale ATCC 23714, Mycoplasma pirum ATCC 106

25960, Mycoplasma pneumoniae NCTC 10119 and Mycoplasma salivarium ATCC 23064 (Public 107

Health England, UK); Streptococcus pneumoniae ATCC 49619, Klebsiella spp. ATCC 700603, 108

Staphylococcus aureus NCTC 6571, Escherichia coli NCTC 10418, Pseudomonas aeruginosa 109

NCTC 10662, Haemophilus influenza NCTC 11931, Legionella pneumophila NCTC 11192, 110

Neisseria gonorrhoeae NCTC 12700 and Mycobacterium tuberculosis H37Rv ATCC 27294 111

(Department of Microbiology, Royal Free NHS Trust, UK); coagulase-negative Staphylococcus, 112

meticillin resistant Staphylococcus aureus, Moraxella catarrhalis, Neisseria meningitidis, 113

Bordetella pertussis, Streptococcus pyogenes, Acinetobacter spp, Corynebacterium spp, Proteus 114

mirabilis and Candida albicans clinical isolates (Department of Microbiology, Royal Free London 115

NHS Foundation Trust); Pneumocystis jirovecii clinical isolate (Microbiology Department, 116

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

6

Raigmore Hospital, UK); Pneumocystic jirovecii (RIE SVC, Edinburgh); Candida spp. (RIE SVC, 117

Edinburgh); Aspergillus fumigatus (RIE SVC, Edinburgh); Chlamydia pneumoniae SA2f (Clinical 118

Microbiology Department, University College London Hospitals NHS Foundation Trust, UK); and 119

viruses (all from RIE SVC, Edinburgh): influenza A, influenza B, RSV, parainfluenza (PIV1-4), 120

human metapneumovirus, human rhinovirus, human coronavirus (hCoV – 229E, OC43, NL63, 121

HKU1, hEV), measles, mumps and human bocavirus (hBoV – 1-4). 122

123

124 Culture 125

Respiratory samples from patients with PAD were inoculated immediately on Mycoplasma 126

Experience agar (Mycoplasma Experience Ltd, Reigate, UK) and incubated at 36°C in gas jars 127

containing CO2 gas packs (Oxoid, Basingstoke, UK). A small number of cultures that had been 128

stored at 4°C for less than 4 days were included as this has been shown previously not to affect the 129

viability of M. amphoriforme (data not shown). Potential M. amphoriforme colonies were detected 130

microscopically with a 40× magnification and their identity was confirmed by the M. amphoriforme 131

16S PCR and sequencing. Primary cultures contaminated with other microorganisms were re-132

cultured using Sputasol treated samples that had been stored at -20°C. Mycoplasma culture was also 133

performed on 16S PCR positive samples from the RFL Chest Clinic, but was not performed on 134

samples from patients with LRTI in general practice as these had been heat killed prior to storage at 135

–70°C and was not performed on RIE respiratory samples submitted for viral diagnosis. 136

137

Extraction of DNA 138

DNA was extracted from control organisms using the Wizard Genomic DNA extraction kit 139

(Promega, Southampton, UK) following manufacturer’s instructions using the protocol for Gram 140

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

7

negative bacteria. Extraction of DNA from sputum and throat swab samples was performed using a 141

Chelex based extraction method: following centrifugation at 13000 × g for 10 min, resulting pellets 142

were washed three times with sterile phosphate buffered saline (PBS), resuspended with PCR grade 143

water (5× the pellet volume) and vortexed with 10 % Chelex (Sigma, Poole, UK) at a ratio of 1:1. 144

After incubation at 56°C for 30 min followed by 94°C for 5 min the samples were vortexed, cooled 145

on ice and then centrifuged at 13000 × g for 2 min with the resulting supernatants being used for 146

PCR. 147

The DNA from respiratory samples for respiratory virus screening was extracted using the 148

easyMAG (bioMérieux, UK) and eluted into 100 μl volume. All extracts were stored at -20°C until 149

used. 150

151

M. amphoriforme 16S PCR 152

All oligonucleotides used in this study are listed in Table 1. The DNA extracts from all patient 153

groups were tested for the presence of M. amphoriforme by the 16S PCR as previously described 154

(2). The identity of the amplicons from at least the first positive sample from each patient was 155

confirmed by sequencing using standard Sanger sequencing protocols. The sequences were analysed 156

using BioNumerics software version 3.5 (Applied Maths) and aligned using the CLUSTAL W 157

multiple sequence alignment programme (8). The consensus sequences were compared to the 16S 158

rDNA sequence obtained from the preliminary contiguous M. amphoriforme strain A39T whole 159

genome sequence obtained from Wellcome Trust Sanger Institute. 160

161

M. amphoriforme qPCRs 162

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

8

The oligonucleotides for the M. amphoriforme udg (Table 1) quantitative real-time PCR were 163

designed and optimised. The optimised M. amphoriforme qPCR protocol consisted of 5μl template 164

DNA, 1x Invitrogen Platinum® QPCR SuperMix-UDG (Invitrogen, UK), 7 mM MgCl2, 0.3 µM 165

primer MAudgF, 0.9 µM primer MAudgR and 0.25 µM probe MAudgP in a final volume of 25 µl. 166

The reactions were performed in a Rotor-Gene 3000 (Qiagen, UK) with cycling conditions of 95°C 167

for 3 min, followed by 35 cycles of 95°C for 15 s and 58°C for 60 s. Results were analysed with the 168

cycle threshold set at 0.03. The standard curves were constructed in two independent experiments on 169

serial ten-fold dilutions of M. amphoriforme DNA in triplicates. The specificity of the assay was 170

confirmed by the amplification of 1 ng of DNA from control organisms listed above in duplicate. 171

The identity of amplicons was confirmed by Sanger sequencing. 172

The udg qPCR was performed on the DNA extracts from patients with PAD. Samples were tested 173

neat, diluted 1/10 and spiked (4 μl of sample and 1 μl pg/μl M. amphoriforme DNA, to detect 174

sample inhibition). Samples positive for both the neat and ten-fold DNA dilution were considered 175

positive, samples that were only positive for either the neat or the ten-fold dilution were considered 176

equivocal. To avoid bias due to sample storage all samples with discrepant results were retested 177

using the 16S PCR. 178

179

A quantitative real-time PCR assay targeting variable region of 23S rDNA unique for M. 180

amphoriforme was designed and optimised. Aliquots of 2 μl template DNA were amplified in a 20 181

μl reaction, using 1× Sso Fast mix (Bio-Rad, UK) and 200 nM of each primer (Table 1). The PCR 182

reactions were carried out in a RotorGene Q thermocycler (Qiagen, UK) set to thermal cycling 183

programme of 95°C, 2 min; 40 cycles of 95°C, 15 s and 60°C, 1 min, and fluorescence detection at 184

(λexc=470 nm, λem=510 nm); and a final melt curve analysis. The specificity of the assay was tested 185

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

9

in silico and in vitro by amplification of non-M. amphoriforme DNA. The detection and 186

quantification limits of the assay were established on M. amphoriforme NCTC 11740 DNA. To 187

further confirm the specificity of the assays the identity of the amplicons was confirmed by 188

sequencing. 189

The 23S PCR was used for M. amphoriforme identification in RIE respiratory samples for viral 190

screening. The DNA was pooled in groups of ten. The DNA from individual samples from positive 191

pools underwent the same 23S PCR amplification to determine individual results. 192

193

194

RESULTS 195

196

Analytical specificity and sensitivity of M. amphoriforme-specific qPCR assays 197

We designed two novel qPCR assays for the identification of M. amphoriforme. Both assays were 198

screened for the specificity in silico and experimentally tested against the DNA of 35 isolates 199

including Mycoplasma spp. and respiratory pathogens. Both qPCR assays were positive only for M. 200

amphoriforme. PCR products were, however, obtained for A. laidlawi, M. alvi and M. genitalium 201

with the 16S PCR. The limit of quantification was 0.01 – 0.1 pg of M. amphoriforme DNA, 202

equivalent to 9-90 organisms per reaction for the udg PCR and 20.6 [2.8 to 149.2, 95% CI] copies 203

per reaction for the 23S PCR, respectively. 204

205

M. amphoriforme in patients attending Immunodeficiency Clinic 206

M. amphoriforme culture, 16S PCR and the udg qPCR were performed on 281 sputum samples from 207

88 patients with PAD attending the RFL Immunodeficiency Clinic. Of these, culture was performed 208

on 278, 16S PCR on 275 and udg qPCR on 263 samples, respectively. M. amphoriforme was 209

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

10

detected by culture and/or PCR in at least one sample of 17 (19.3%) patients. A positive culture was 210

obtained for 10 patients (37 samples of 278, 13.3%), 16S PCR was positive for 17 patients (70/275, 211

25.5%) and qPCR for 14 patients (64/263, 24.3%). The results are summarized in Table 2. The 212

GenBank accession numbers for M. amphoriforme 16S rDNA from this patient group are 213

HM235425 to HM235439. Multiple samples tested positive for 11 of the 13 patients where multiple 214

samples were received with positivity lasting for between 197 and 1627 days. Estimated bacterial 215

loads were ≥ 106 organisms per ml of sputum in at least one sample for 10 positive patients. Routine 216

microbiology results were available for 70 samples from 15 M. amphoriforme positive patients and 217

for 39 samples from 18 matched negative patients. Known respiratory pathogens including H. 218

influenzae, S. pneumoniae, M. catarrhalis and P. aeruginosa were found in more M. amphoriforme 219

negative sputum samples (59%) compared with M. amphoriforme positive samples (24%). H. 220

influenzae was the most commonly isolated pathogen (18% of all samples), and was found 221

significantly less often in M. amphoriforme positive samples (Fisher’s exact test, p=0.003). 222

223

M. amphoriforme in patients attending Chest Clinic 224

A total of 38 sputum samples from 37 patients attending the RFL Chest Clinic were tested. Of the 225

patients, 17 had a diagnosis of chronic obstructive pulmonary disease, 14 had bronchiectasis and one 226

patient had both conditions. Culture results indicated normal respiratory tract flora in 15, H. 227

influenzae in four cases, four samples with S. aureus and samples with single isolates of 228

Acinetobacter, Pseudomonas and Citrobacter spp. In this group, there was a single sample positive 229

for M. amphoriforme (Table 2); a patient who had been taking clarithromycin for an exacerbation of 230

symptoms and in whom no other significant pathogen was found. The GenBank accession number 231

for the M. amphoriforme 16S sequence from this sample is HM235449. 232

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

11

233

M. amphoriforme in patients with suspected LRTI 234

M. amphoriforme was detected in four (5%) out of 80 patients with LRTI recruited from general 235

practices (1/80 throat swabs, 3/50 sputum samples) by 16S PCR (Table 2). The identity of all PCR 236

positive amplicons was confirmed by sequencing (GenBank accession numbers HM235442 to 237

HM235446). None of the control samples (49 throat swabs from healthy individuals) were positive. 238

All four M. amphoriforme positive samples were samples taken from patients with clinical signs of 239

acute LRTI, including raised pulse rates, respiratory rates and CRP concentration compared with the 240

controls. None had a history of recent travel, alcohol consumption or steroid treatment. Other 241

respiratory organisms were detected in two of the M. amphoriforme positive patients; coronavirus, 242

human rhinovirus, H. influenzae plus Streptococcus pneumoniae in the patient with a positive throat 243

swab and enterovirus in a patient with positive sputum. 244

245

M. amphoriforme in patients with suspected respiratory viral infection 246

The respiratory samples screened for suspected LRTI used in this study were collected between 1st 247

March 2011 and 11th March 2012. Out of 10747 (3496 adults and 7251 children) respiratory samples 248

from 7139 patients (2524 adults and 4615 children), 23 samples from 19 patients (6 adults, 13 249

children) tested positive by M. amphoriforme 23S qPCR (Table 2). The positive samples involved 250

nasopharyngeal secretions (6), nose throat swab (1), throat swabs (8), throat swabs for virology (3), 251

sputum (1) and induced sputum (1), and they originated from Accident and Emergency (12), 252

Intensive Treatment Unit / High Dependency Unit (4), Children’s Ward (4), Infectious Diseases (1), 253

Haematology (1) and Neonatal Unit (1). No other respiratory pathogen was found in 13 samples and 254

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

12

10 samples had viral co-infection as detected by real-time PCR: rhinovirus (4), RSV (3), adenovirus 255

and influenza B (1), human metapneumovirus (hMPV) (1) and parainfluenza virus 2 (1). 256

257

258

DISCUSSION 259

To better understand the epidemiology and pathogenesis of M. amphoriforme infection it is 260

necessary to develop new sensitive and quantitative tools for diagnosis. Due to the fastidious growth 261

of human mycoplasmas, sensitive molecular tools are an essential pre-requisite for their 262

identification in order to diagnose infection in a timely manner so that antimicrobial treatment can 263

be initiated. In this paper we describe two real-time PCR assays, define their specificity and evaluate 264

them in a clinical practice environment. 265

266

Specificity of the qPCR assays 267

The qPCR assays target udg gene and M. amphoriforme-specific region of 23S rDNA. Both qPCRs 268

were specific for M. amphoriforme, being negative for all other tested species. In contrast, the 16S 269

PCR was positive for three mycoplasma related species: A. laidlawii, M. genitalium and M. alvi, 270

respectively. A. laidlawii can be found in the human oropharynx and although M. genitalium is 271

primarily a genitourinary tract pathogen of humans, there have been reports of its detection from 272

respiratory samples (9); they may, therefore represent a risk of false positive results. M. alvi has only 273

been found in cattle and there is no evidence of its presence in humans (10). The high specificity of 274

the real time assays was further confirmed by sequencing of products which showed that all positive 275

samples contained M. amphoriforme specific sequence. 276

277

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

13

The assays are able to detect an estimated single to several copies per reaction and can be used to 278

measure the bacterial load. High sensitivity of the qPCR reactions may be important in defining the 279

pathogenic potential of M. amphoriforme in future studies as has been the case for other organisms 280

such as M. genitalium (11, 12). Moreover, a sensitive detection method will improve detection if 281

sub-optimal samples are used, as it is not yet clear what is the primary niche of M. amphoriforme in 282

the human host. 283

284

M. amphoriforme in samples from patients with immunodeficiency 285

The 16S PCR and qPCR provide more sensitive detection than culture, identifying M. amphoriforme 286

in 17 patients (25.5% positive samples) and 14 patients (24.3% positive samples), respectively, 287

versus 10 culture-positive patients (13.3% positive samples). The qPCR gave an equivocal signal for 288

one sample and was negative for another sample for two patients with only a single sample available 289

for the analysis. However, these samples were positive by the 16S PCR but negative by culture. 290

These results may have arisen through undetected inhibition or loss of DNA during extraction. 291

There was a single sample from one patient positive by the 16S PCR that was not available for the 292

qPCR and showed negative by culture. The high incidence of M. amphoriforme (19.3%) in sputa of 293

PAD patients suggests that it may be an important cause of infection in this patient group. Although 294

it is difficult to assign the clinical significance of M. amphoriforme in this complex group of 295

patients, our data show that M. amphoriforme can infect PAD patients chronically and may 296

contribute to LRTI and the pathogenesis of lung disease. Further research should be conducted to 297

characterise M. amphoriforme pathogenicity in this patient group. 298

299

M. amphoriforme from samples from the Chest Clinic and from patients with LRTI 300

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

14

A single sample from 37 patients attending the Chest Clinic was positive for M. amphoriforme. 301

These patients are known to be susceptible to a wide range of pathogens causing chronic sepsis and 302

further studies in larger groups of patients are required. The detection rate of M. amphoriforme (5%) 303

in patients with acute signs of LRTI recruited from general practices was similar to that of other 304

known respiratory pathogens, including Haemophilus influenzae (6%), coronaviruses (6%) and 305

parainfluenza viruses (4%) (5). Co-infections were a common feature of this patient group (22.5% of 306

patients and 4% of controls) and, therefore, the co-infection of the M. amphoriforme positive sample 307

with other organisms does not exclude its aetiological role in LRTI. It was notable that M. 308

amphoriforme was not detected in control subjects as these samples were exclusively throat swabs. 309

It opens the possibility that this observation is due to the sample type or that M. amphoriforme is a 310

primary respiratory pathogen. However, one throat swab from a patient was M. amphoriforme 311

positive in this study and throat swabs are recommended for the detection of other Mycoplasma spp. 312

(13). In addition, other respiratory pathogens such as S. pneumoniae and viral pathogens were 313

detected using these throat swabs at their expected frequency (5). 314

315

M. amphoriforme in samples from patients with suspected respiratory viral infection 316

The largest group in this study was a one-year collection of 10747 respiratory samples from patients 317

with suspected respiratory infection submitted for virological testing. The infection with M. 318

amphoriforme was found to be uncommon within this group with the incidence of 0.21%. 319

The low incidence is not likely to be caused by pooling as this approach has previously proved 320

successful in the detection of hMPV in clinical samples (14). The study described here is a pilot 321

using sampling protocols and DNA extraction methods that are not yet optimised for this organism. 322

Thus, the low detection rate may be because this is not an optimal sample. Additionally, other 323

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

15

Mycoplasma species have periodicity in their detection rate, for example M. pneumoniae infection 324

increases in prevalence every four to seven years (15, 16) The longest study period reported here 325

was one year, thus, longitudinal studies are now required to elucidate the periodicity of M. 326

amphoriforme infection. In this study, positive results were mostly found in children (68% of the 327

positive patients) but this may reflect the distribution of the samples submitted for testing. An age 328

cross-sectional study is now required. Viral co-infection was present in 10 M. amphoriforme 329

positive samples, all from children. Interestingly, viral infection was not detected in any of the M. 330

amphoriforme positive samples from adult patients. The results from this preliminary study will 331

provide the basis for a larger study in a wide range of samples from patients presenting with 332

symptoms and signs of LRTI. 333

334

The results reported here are important pilot data for M. amphoriforme and the first step in 335

understanding its wider pathogenicity. Taken together these data provide support for the idea that M. 336

amphoriforme may be a primary respiratory pathogen. Despite this, these studies should be repeated 337

by other groups in different countries and we are currently working with partners to perform such 338

work. The importance of this paper is that it provides the methodology to assist other groups in 339

diagnosing M. amphoriforme and it is only by increasing the number of patients identified with this 340

organism that we will be able to determine its pathogenic potential with certainty. 341

342

343

ACKNOWLEDGEMENTS 344

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

16

This work was supported by a Peter Samuel Royal Free Fund Grant; the Primary Immunodeficiency 345

Association, the Special Trustees of the Royal Free London NHS Foundation Trust, Hampstead, and 346

the University of St Andrews Medical School. 347

348

349

Table1 Oligonucleotides used in this study. 350

351

Target Oligonucleotide (5’-3’) Product size (bp)

udg MAudgF TGCGGCCGATAAAACCGAAATAT

92 MAudgR TTTCGAAAAGGGTTTGCTACCAA Probe FAM-TTGTGCTCATCCTTCACCCTTTAGTGTGCA-BHQ1

23S rDNA Forward GGGGTTCAAATAACAAGTC

106 Reverse CGTGATATATGGCTCTTCG

16S rDNA Amph-f: AAGCTAGTAAAGGAAATGTTATT Amph-r: ACTATAGAAATATAGTC

594

352

Table 2 M. amphoriforme positive rates for different patient groups tested by culture, 16S PCR and 353

qPCR. 354

355

Patient Group Culture 16S rDNA PCR qPCR Total Positives

Immunodeficiency Clinic

11.36% (10/88) 19.32% (17/88) 16.09% (14/87)

19.32% (17/88)

Chest Clinic NA 2.70% (1/37) NA 2.70% (1/37)

LRTI patients NA 3.08% (4/80) NA 3.08% (4/80)

Suspected viral LRTI NA NA 0.27% (19/7139)

0.27% (19/7139)

356

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

17

357

REFERENCES 358

359 1. Webster D, Windsor H, Ling C, Windsor D, Pitcher D. 2003. Chronic Bronchitis in 360

Immunocompromised Patients: Association with a Novel Mycoplasma Species. European 361 Journal of Clinical Microbiology & Infectious Diseases 22:530–534. 362

2. Pitcher DG. 2005. Mycoplasma amphoriforme sp. nov., isolated from a patient with chronic 363 bronchopneumonia. INTERNATIONAL JOURNAL OF SYSTEMATIC AND 364 EVOLUTIONARY MICROBIOLOGY 55:2589–2594. 365

3. Pereyre S, Renaudin H, Touati A, Charron A, Peuchant O, Hassen AB, Bébéar C, 366 Bébéar CM. 2009. Detection and susceptibility testing of Mycoplasma amphoriformeisolates 367 from patients with respiratory tract infections. Clinical Microbiology and Infection. 368

4. Hatchel JM. 2006. Ultrastructure and gliding motility of Mycoplasma amphoriforme, a 369 possible human respiratory pathogen. Microbiology (Reading, Engl.) 152:2181–2189. 370

5. Creer DD, Dilworth JP, Gillespie SH, Johnston AR, Johnston SL, Ling C, Patel S, 371 Sanderson G, Wallace PG, McHugh TD. 2006. Aetiological role of viral and bacterial 372 infections in acute adult lower respiratory tract infection (LRTI) in primary care. Thorax 373 61:75–79. 374

6. Woodhead M, Blasi F, Ewig S, Garau J, Huchon G, Ieven M, Ortqvist A, Schaberg T, 375 Torres A, van der Heijden G, Read R, Verheij TJM, Joint Taskforce of the European 376 Respiratory Society and European Society for Clinical Microbiology and Infectious 377 Diseases. 2011. Guidelines for the management of adult lower respiratory tract infections - 378 Summary. Clinical Microbiology and Infection 17:1–24. 379

7. NICE. 2009. Respiratory tract infections – antibiotic prescribing 1–121. 380 8. Chenna R, Sugawara H, Koike T, Lopez R, Gibson TJ, Higgins DG, Thompson JD. 381

2003. Multiple sequence alignment with the Clustal series of programs. Nucleic Acids Res. 382 31:3497–3500. 383

9. Baseman JB, Dallo SF, Tully JG, Rose DL. 1988. Isolation and characterization of 384 Mycoplasma genitalium strains from the human respiratory tract. Journal of Clinical 385 Microbiology 26:2266–2269. 386

10. Pettersson B, Uhlén M, Johansson KE. 1996. Phylogeny of some mycoplasmas from 387 ruminants based on 16S rRNA sequences and definition of a new cluster within the hominis 388 group. Int. J. Syst. Bacteriol. 46:1093–1098. 389

11. Taylor-Robinson D, Jensen JS. 2011. Mycoplasma genitalium: from Chrysalis to 390 multicolored butterfly. Clinical Microbiology Reviews 24:498–514. 391

12. Edberg A, Jurstrand M, Johansson E, Wikander E, Höög A, Ahlqvist T, Falk L, Jensen 392 JS, Fredlund H. 2008. A comparative study of three different PCR assays for detection of 393 Mycoplasma genitalium in urogenital specimens from men and women. Journal of Medical 394 Microbiology 57:304–309. 395

13. Dorigo-Zetsma JW, Zaat SA, Wertheim-van Dillen PM, Spanjaard L, Rijntjes J, van 396 Waveren G, Jensen JS, Angulo AF, Dankert J. 1999. Comparison of PCR, culture, and 397 serological tests for diagnosis of Mycoplasma pneumoniae respiratory tract infection in 398 children. Journal of Clinical Microbiology 37:14–17. 399

14. Manning A, Russell V, Eastick K, Leadbetter GH, Hallam N, Templeton K, Simmonds 400

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from

18

P. 2006. Epidemiological profile and clinical associations of human bocavirus and other 401 human parvoviruses. J. Infect. Dis. 194:1283–1290. 402

15. Bébéar C, Pereyre S, Bébéar CM. 2006. Principles and Practice of Clinical Bacteriology, 403 pp. 305–315. In Gillespie, SH, Hawkey, PM (eds.), Second Edition. John Wiley & Sons, Ltd, 404 Chichester, UK. 405

16. Jacobs E. 2012. Mycoplasma pneumoniae: now in the focus of clinicians and 406 epidemiologists. Euro Surveill. 17. 407

408

on March 21, 2018 by guest

http://jcm.asm

.org/D

ownloaded from