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Susceptibility testing of sequential isolates of Aspergillus fumigatus recovered from treated patients

Eric Dannaoui¹, Joseph Meletiadis², Anna-Maria Tortorano³, Françoise Symoens⁴, Nicole Nolard⁴, Maria-Anna Viviani³, Marie-Antoinette Piens¹, Bernadette Lebeau⁵, Paul E. Verweij² and Renée Grillot⁵ and the EBGA Network

¹Laboratoire de Parasitologie, Mycologie Médicale et Pathologie Exotique, Université Claude Bernard Lyon I, 69373 Lyon Cedex 08, France ²Department of Medical Microbiology, University Medical Center St Radboud, 6500 HB Nijmegen, The Netherlands ³Istituto di Igiene e Medicina Preventiva, Università degli Studi –IRCCS Ospedale Maggiore, 20122 Milan, Italy ⁴Scientific Institute of Public Health, B-1050 Brussels, Belgium ⁵Laboratoire Interactions Cellulaires Parasite–Hôte, Université Joseph Fourier Grenoble I, 38706 Grenoble Cedex, France ⁶European Research Group on Biotypes and Genotypes of Aspergillus fumigatus

Correspondence Eric Dannaoui dannaoui{at}pasteur.fr

Received May 28, 2003
Accepted November 6, 2003

Two-hundred sequential Aspergillus fumigatus isolates recovered from 26 immunocompromised patients with invasive aspergillosis or bronchial colonization were tested for their in vitro susceptibility to posaconazole, itraconazole, voriconazole, terbinafine and amphotericin B. Twenty-one patients were treated with amphotericin B and/or itraconazole. Antifungal susceptibilities of the isolates recovered before treatment were not significantly different from those of isolates recovered after the onset of antifungal therapy. The highest MICs were 0.125, 0.5, 0.5, 1 and 1 µg ml^-1 for posaconazole, itraconazole, voriconazole, terbinafine and amphotericin B, respectively. It is concluded that the emergence of resistance in A. fumigatus during antifungal therapy with amphotericin B or itraconazole is an uncommon phenomenon.

	INTRODUCTION

TOP INTRODUCTION METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Invasive infections caused by Aspergillus fumigatus are an increasing problem in immunocompromised patients, in particular those with neutropenia and in transplant recipients (Denning 1998; Mehrad et al., 2001). In these patients, invasive aspergillosis is associated with high mortality rates.

Amphotericin B is the drug of choice for treatment of invasive aspergillosis, but the overall success rate remains low (Denning et al., 1998) and its use as a therapeutic agent is limited by its toxicity (Georgopapadakou & Walsh, 1994). Itraconazole is significantly better tolerated than amphotericin B and it has been successfully used to treat patients with invasive aspergillosis (Denning, 1998).

More recently, new drugs with anti-Aspergillus activity, including azoles and echinocandins, have been evaluated for their use as therapeutic agents. Voriconazole has been shown to be efficacious in treating invasive aspergillosis in patients (Denning et al., 2002) and preliminary data have shown that posaconazole was efficacious as salvage treatment of invasive aspergillosis in patients refractory to, or intolerant of, standard antifungal therapies (Hachem et al., 2000). Caspofungin is also effective for the treatment of invasive aspergillosis and this drug has a good safety profile (Deresinski & Stevens, 2003).

The extensive use of azole compounds such as fluconazole in the treatment of infections has been associated with the emergence of antifungal-resistant Candida strains, particularly in HIV-infected patients (Sanglard & Odds, 2002). In vitro resistance of A. fumigatus strains to itraconazole has been described (Chryssanthou, 1997; Dannaoui et al., 1999b, 1997; Verweij et al., 2002) and for some of these strains the resistance has been confirmed in vivo in animal models of aspergillosis (Dannaoui et al., 1999a, 2001; Denning et al., 1997). Itraconazole resistance in A. fumigatus has been reported both before exposure to the drug (Dannaoui et al., 1999a; Denning et al., 1997) and after antifungal therapy (Chryssanthou, 1997; Dannaoui et al., 2001; Denning et al., 1997; Verweij et al., 2002).

A comprehensive review of the recent literature has shown that about 2.1 % of more than 900 strains of A. fumigatus were resistant to itraconazole (Moore et al., 2000). Nevertheless, the frequency of acquired resistance under long-term therapy among A. fumigatus isolates is largely unknown.

In vivo resistance to amphotericin B has been reported in animal models (Johnson et al., 2000; Verweij et al., 1998a), as well as in neutropenic patients with invasive aspergillosis (Lass-Florl et al., 1998).

In this study, performed within the European Research Group on Biotypes and Genotypes of Aspergillus fumigatus (EBGA Network), 26 patients with A. fumigatus colonization or infection were submitted to a prolonged clinical and mycological follow-up at three European hospital centres. Sequential isolates obtained before and after the onset of antifungal therapy were tested for their susceptibility to five antifungal drugs.

	METHODS

TOP INTRODUCTION METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Patients and isolates.

Two-hundred clinical isolates of A. fumigatus from 26 patients were included in the study. These strains were isolated at three European hospital centres [Centre A, Lyon, France (16 patients, 127 isolates); Centre B, Grenoble, France (6 patients, 26 isolates); Centre C, Milan, Italy (4 patients, 47 isolates)] from immunocompromised patients with invasive aspergillosis or lung transplant recipients with bronchial colonization.

The 200 isolates were recovered from 114 respiratory samples, which were mostly broncho-alveolar lavages. For 21 samples (representing 107 isolates obtained from 12 different patients), different colonies were isolated from the same sample. Table 1 summarizes the clinical and therapeutic data for all 26 patients. Sixteen patients had invasive aspergillosis and 10 had bronchopulmonary colonization after lung transplant. For the 21 patients who received antifungal therapy, 11 were treated with amphotericin B alone, eight received amphotericin B and itraconazole, and two were treated with itraconazole alone (Table 1).

The strains were identified in each centre by conventional techniques based on classical morphological criteria (Raper & Fennell, 1965). Following their isolation, the strains were sent to the Institute of Hygiene and Epidemiology Mycology (IHEM, Brussels, Belgium), where the collection was maintained.

Antifungal susceptibility testing The antifungal susceptibility testing done following an NCCLS-based methodology (NCCLS, 1998) was performed in three centres: Centre A (76 isolates), Centre C (65 isolates) and Centre D, Nijmegen, The Netherlands (59 isolates). Previous studies had shown that good intra- and inter-laboratory reproducibility was obtained in these centres (Tortorano et al., 2000, 2002). All the serial isolates from the same patient were tested in one centre. According to the protocol, resistant isolates would have been re-tested in the other centres.

Three reference strains, Candida krusei ATCC 6258, Candida parapsilosis ATCC 22019 and itraconazole-resistant A. fumigatus IHEM 13936 (NCPF 7100; Denning et al., 1997), were included in the study to ensure quality control.

Medium.

RPMI 1640 medium with L-glutamine but without sodium bicarbonate (Sigma Chemical) buffered to pH 7.0 with 0.165 M MOPS (Sigma Chemical) was used as the test medium. The same batch of medium was used in the three centres.

Inoculum preparation.

The lyophilized isolates were sent from the IHEM culture collection to the three participating centres, where they were reconstituted with sterile distilled water. Isolates were subcultured twice on potato dextrose agar (PDA) to ensure viability and were then grown on PDA slants for 5 days at 35 °C. Stock spore suspensions were prepared by washing the surface of the slants with 1.5 ml of sterile saline containing 0.05 % Tween 80. The spore suspensions were transferred to a sterile tube, turbidity was measured spectrophotometrically at 530 nm and adjusted to an optical density ranging from 0.09 to 0.11 (80–82 % transmittance), to yield an initial inoculum of 5 x 10⁵ to 5 x 10⁶ c.f.u. ml^-1 (Espinel-Ingroff et al., 1997). Each suspension was diluted 1 : 50 (v/v) in the medium to obtain twice the desired inoculum. Inoculum size for each isolate was checked by quantitative colony counts on Sabouraud agar plates.

Broth microdilution test.

MICs were determined by a broth microdilution technique using sterile microdilution plates (96 U-shaped wells). The drugs tested included posaconazole (SCH56592; Schering-Plough Research Institute), itraconazole (Janssen Pharmaceutica), voriconazole (Pfizer Central Research), terbinafine (Novartis Pharma) and amphotericin B (Sigma Chemical). The same batches of antifungal drugs were used in the three centres. Drugs were dissolved at a concentration of 1600 µg ml^-1 in DMSO. Drug dilutions were prepared as twice the strength of the final concentration by following the additive two-fold drug dilution NCCLS scheme (NCCLS, 1998). Final concentrations of the antifungal agents were 0.03–16 µg ml^-1. For each strain, a growth control well containing medium plus 0.5 % of the corresponding solvent was included. Microdilution trays were kept at -20 °C until the day of testing.

Incubation and MIC determination.

On the day of the test, each well of the microtitre plates containing 100 µl of the diluted drug concentrations was inoculated with 100 µl of the inoculum suspension. Microplates were incubated at 35 °C and MICs were determined visually after 48 h incubation. MIC determinations were done in duplicate in two independent experiments.

Visual MIC determination.

Microplates were read visually with the aid of a microtitre reading mirror, and the growth in each well was compared to that of the growth control. Each well was then given a numerical score according to the NCCLS guidelines (NCCLS, 1998): 4, no reduction in growth; 3, 25 % reduction in growth; 2, 50 % reduction in growth; 1, 75 % or more reduction in growth; 0, no growth (optically clear). MIC end-points were defined as the lowest drug concentration that had a score of 0 for amphotericin B and a score of 2 for the other drugs.

Data analyses.

MIC ranges and MICs for 50 % (MIC₅₀s) and 90 % (MIC₉₀s) of the isolates tested were determined. For calculations, the low off-scale MICs were left unchanged. Differences in the distributions of MICs were determined by the Kruskal–Wallis test or the Friedman test as appropriate. Statistical analyses were performed using GRAPHPAD PRISM version 3.00 for WINDOWS (GRAPHPAD Software). Statistical significance was defined as P 0.05.

	RESULTS

TOP INTRODUCTION METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
The median elapsed time between the first and last isolate was 49 days (range, 2–1079 days). The median number of isolates obtained per patient was six (range, 2–37) and the median number of isolates exposed to treatment was two per patient (range, 0–37). The maximum exposure time to the drug before isolation of A. fumigatus was 82 and 401 days for amphotericin B and itraconazole, respectively (Table 1).

Table 2 summarizes the in vitro activity of the five antifungal drugs against the 200 isolates of A. fumigatus, as determined after 48 h incubation. Overall, posaconazole was the most active drug (P < 0.001) against A. fumigatus, with terbinafine having the widest range of MICs. The highest MICs were 0.125, 0.5, 0.5, 1 and 1 µg ml^-1 for posaconazole, itraconazole, voriconazole, terbinafine and amphotericin B, respectively.

For 21 of the specimens, several colonies from the same specimen were tested: in all cases, MICs for itraconazole, voriconazole and posaconazole were within three log2 dilutions; a difference of four dilutions was noted for amphotericin B in one sample and for terbinafine in another sample (data not shown).

Table 3 summarizes the antifungal susceptibility of pre- and post-treatment isolates (i.e. isolates recovered before or after the onset of the treatment, respectively) to amphotericin B and itraconazole. One-hundred post-treatment isolates were recovered from the 16 patients treated with amphotericin B. Amphotericin B MICs for these isolates was not significantly different from MICs for the 91 pre-treatment isolates. In the same way, MICs for itraconazole were similar for the 67 isolates (from 10 patients) recovered following itraconazole therapy and for the 131 pre-treatment isolates.

	DISCUSSION

TOP INTRODUCTION METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Emergence of antifungal resistance in Candida albicans associated with treatment failures has been well documented in AIDS patients with oropharyngeal candidiasis (Sanglard & Odds, 2002). However, few studies have been conducted to evaluate the potential emergence of antifungal resistance in patients with infections due to filamentous fungi.

In vitro itraconazole resistance in A. fumigatus has been reported (Chryssanthou, 1997; Dannaoui et al., 1999b; Denning et al., 1997; Verweij et al., 2002) and this resistance was confirmed for some strains in animal models of aspergillosis (Dannaoui et al., 1999a, 2001; Denning et al., 1997), suggesting a clinical significance for these in vitro results. Both primary and acquired resistance have been documented. In some cases, itraconazole-resistant strains have been isolated before any exposure to the drug (Dannaoui et al., 1999b; Denning et al., 1997). Development of resistance in patients during itraconazole therapy has also been reported in some studies. In one study, a strain was isolated during treatment with itraconazole but was genotypically distinct from the initial strain cultured before treatment (Denning et al., 1997). Chryssanthou (1997) reported four itraconazole-resistant isolates obtained from three patients after 5 months to 3 years of itraconazole therapy. Nevertheless, as the genotypic profiles of these strains were not studied, it was not possible to determine whether the strains acquired resistance or if the patients were subsequently infected by another strain from the environment (Verweij et al., 1998b). In a large study in The Netherlands, three A. fumigatus isolates resistant to itraconazole were recovered from a lung transplant recipient after several months of itraconazole therapy (Verweij et al., 2002). In another study, acquisition of itraconazole resistance after 4–5 months of therapy with this antifungal drug was documented (Dannaoui et al., 2001). However, the frequency of this phenomenon in patients undergoing long-term therapy with itraconazole is unknown.

In the present study, A. fumigatus isolates sequentially obtained from patients with chronic pulmonary colonization or invasive aspergillosis were tested for their antifungal susceptibilities, and emergence of resistance to itraconazole was not detected. Nevertheless, it should be noted that only five patients received itraconazole therapy for more than 2 months.

All the isolates exhibited low amphotericin B MICs, of 1 mg l^-1, and there were no differences between isolates exposed to this drug and isolates recovered without treatment. This is in accord with a previous study in which Aspergillus isolates recovered from six patients receiving amphotericin B for invasive aspergillosis were tested for their antifungal susceptibility (Moosa et al., 2002). In that study, emergence of resistance during therapy was also not detected. However, the amphotericin B resistance of Aspergillus spp. is difficult to detect in vitro and the correlation between MIC and clinical outcome in animal models is poor (Johnson et al., 2000; Mosquera et al., 2001; Verweij et al., 1998a). Nevertheless, it has been shown that in vitro susceptibility testing of amphotericin B is a good predictor of clinical outcome in neutropenic patients with invasive aspergillosis (Lass-Florl et al., 1998).

It has to be pointed out that the genetic diversity of A. fumigatus is very high, particularly in lung transplant recipients or cystic fibrosis patients with bronchial colonization (Symoens et al., 2001; Verweij et al., 1996). More than 40 strains (isolated from eight lung transplant recipients) used in the present study were previously typed by three molecular methods (Symoens et al., 2001). The results showed that the same patient could be sequentially colonized by different genotypes. This could explain in part the low frequency of resistance; one given isolate may be exposed to the drug for only a limited period of time due to the replacement of the isolate by a new one. Another possible explanation is that the hyphal form of the fungus (which is the only stage found in infected tissues) is less susceptible to acquisition and in vivo spread of mutations under drug pressure than the conidiogenous form.

For the other drugs tested in the present study, the highest MICs were 0.125, 0.5 and 1 µg ml^-1 for posaconazole, voriconazole and terbinafine, respectively. Although break points have not been established for moulds, none of the isolates could be considered resistant to any of these three drugs. Overall, posaconazole was the most active of the five antifungal drugs tested. Nevertheless, it has to be pointed out that this will not necessarily be translated in vivo as many other pharmacokinetic considerations have to be taken into account.

In conclusion, the data of the present study suggest that emergence of resistance in A. fumigatus during antifungal therapy is an uncommon phenomenon.

Members of the EBGA Network (European Research Group on Biotypes and Genotypes of Aspergillus fumigatus), European Concerted Action no. BMH4-CT-97-2481, 1997–2000, are as follows. Co-ordinator, R. Grillot. Laboratoire de Parasitologie et Mycologie Médicale, Université Claude Bernard Lyon I (M. A. Piens, E. Dannaoui, M. Perraud & M. F. Monier) et Information Médicale, Lyon, France (F. Chapuis). Laboratoire de Parasitologie-Mycologie Médicale et Moléculaire, Faculté de Médecine, Grenoble, France (R. Grillot, B. Lebeau & J. Burnod). Scientific Institute of Public Health, Brussels, Belgium (N. Nolard, F. Symoens, K. Goens & S. Heinemann). Laboratoire d'Immunologie et Parasitologie, Faculté de Pharmacie, Montpellier, France (J. M. Bastide, M. Mallié, D. Castel, S. Berthou, F. Renaud & T. De Meeus). Instituto de Igiene e Medicina Preventiva, Milan, Italy (M. A. Viviani, A. M. Tortorano, A. L. Rigoni & M. Cogliati). Department of Microbiology, University of Leeds, UK (E. G. Evans, R. Barton, R. Ashbee & V. Hopwood). Department of Medical Microbiology, University Medical Center Nijmegen, Nijmegen, The Netherlands (J. F. Meis, A. Voss, P. E. Verweij & J. P. Donnelly). Laboratoire de Mycologie Fondamentale et Appliquée, Faculté de Pharmacie Lyon, France (J. Villard, A. Couble & A. Casoli). Institut für Medizinische Mikrobiologie, Universität GH Essen, Germany (P. M. Rath & R. Ansorg).

	ACKNOWLEDGEMENTS

TOP INTRODUCTION METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
This work was presented in part at the 14th ISHAM Congress, Buenos Aires, Argentina, 8–11 May 2000.

	REFERENCES

TOP INTRODUCTION METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES