Content uploaded by Dominique Sanglard
Author content
All content in this area was uploaded by Dominique Sanglard on Jul 10, 2015
Content may be subject to copyright.
Danie
`
le Maubon
Ce
´
cile Garnaud
Thierry Calandra
Dominique Sanglard
Muriel Cornet
Resistance of
Candida
spp. to antifungal drugs
in the ICU: where are we now?
Received: 15 May 2014
Accepted: 10 July 2014
! Springer-Verlag Berlin Heidelberg and
ESICM 2014
Take-home message: The emergence of
resistance is a warning signal triggering
improvements in antifungal drug use,
particularly in patients for whom the
potential benefit of treatment is unproven.
Practical proposals to detect and prevent the
risk of clinical failure are (i) accurate
assessments of prior antifungal exposure,
(ii) close clinical monitoring of patients
treated with antifungal drugs, (iii) routine
surveillance of in vitro susceptibility testing
and (iv) development of feasible methods
for rapid detection of mutations.
Electronic supplementary material The
online version of this article (doi:
10.1007/s00134-014-3404-7) contains sup-
plementary material, which is available to
authorized users.
D. Maubon (
)
) ! C. Garnaud ! M. Cornet
Parasitologie-Mycologie, Institut de
Biologie et de Pathologie, CHU de
Grenoble, Grenoble, France
e-mail: dmaubon@chu-grenoble.fr
Tel.: ?33 4 76 76 54 90
D. Maubon ! C. Garnaud ! M. Cornet
Laboratoire TIMC-TheREx, UMR 5525
CNRS-UJF, Universite
´
Grenoble Alpes,
Grenoble, France
T. Calandra
Infectious Diseases Service, Department of
Medicine, Centre Hospitalier Universitaire
Vaudois and University of Lausanne,
Lausanne, Switzerland
D. Sanglard
Institute of Microbiology, Centre
Hospitalier Universitaire Vaudois and
University of Lausanne, Lausanne,
Switzerland
Abstract Current increases in anti-
fungal drug resistance in Candida
spp. and clinical treatment failures are
of concern, as invasive candidiasis is
a significant cause of mortality in
intensive care units (ICUs). This
trend reflects the large and expanding
use of newer broad-spectrum anti-
fungal agents, such as triazoles and
echinocandins. In this review, we
firstly present an overview of the
mechanisms of action of the drugs
and of resistance in pathogenic
yeasts, subsequently focusing on
recent changes in the epidemiology of
antifungal resistance in ICU. Then,
we emphasize the clinical impacts of
these current trends. The emergence
of clinical treatment failures due to
resistant isolates is described. We
also consider the clinical usefulness
of recent advances in the interpreta-
tion of antifungal susceptibility
testing and in molecular detection of
the mutations underlying acquired
resistance. We pay particular atten-
tion to practical issues relating to ICU
patient management, taking into
account the growing threat of anti-
fungal drug resistance.
Keywords Antifungal resistance !
Resistance mechanisms !
Candida resistance !
Intensive care unit !
Clinical resistance !
Microbial resistance
Introduction
Invasive candidiasis is a major threat to intensive care
unit (ICU) patients, causing significant mortality. An
early initiation of antifungal therapy is crucial to improve
the prognosis [1, 2]. However, the performance of current
diagnostic tools for confirming the diagnosis remains
limited. Increasing numbers of ICU patients without
documented candidiasis are therefore receiving prophy-
lactic or empirical antifungal treatments in an attempt to
decrease Candida-related mortality. This strategy has
been encouraged by the introduction of new, better-tol-
erated antifungal drugs, such as triazoles and
echinocandins, leading to stronger selective pressure [3,
4]. Antifungal drug resistance was considered less prob-
lematic in Candida spp. than in other pathogens, but
Intensive Care Med
DOI 10.1007/s00134-014-3404-7
REVIEW
recent increases in resistance to both echinocandins and
azoles have led to clinical failures [5, 6]. This is a matter
of concern because of the limited number of drug classes
targeting different fungal components and because the
number of patients at risk receiving treatment is contin-
ually growing, thus further increasing antifungal drug
pressure.
In this review, we firstly summarize the basis of the
mechanisms of action and resistance concentrating on
recent advances that improve our understanding of
antifungal drugs. Then, we describe the current changes
in the epidemiology of Candida spp. resistance. We
enlighten their consequences for responses to antifungal
treatments and for the optimal choice for empiric, pre-
emptive and targeted strategies in ICU patients. The
clinical relevance of the new developments for labora-
tory antifungal drug testing and for the detection of
resistance-associated mutations is discussed with spe-
cific attention paid to practical approaches, to assess the
risk of clinical treatment failure and to improve its
prevention.
Targets and mechanisms of action of systemic
antifungal drugs
Fungi are more closely related to humans than other
pathogens, such as bacteria, limiting the number of
available antifungal targets. Despite the introduction of a
novel drug class exploiting a new target (echinocandins)
and new azole drugs with a broader spectrum of activity
(voriconazole, posaconazole), the antifungal arsenal still
remains restricted.
Antifungal agents acting on the cell wall and/
or plasma membrane
Echinocandins
Caspofungin, micafungin and anidulafungin block cell
wall synthesis by inhibiting (1,3)-b-
D-glucan synthase,
which catalyses the first step in the elongation of (1,3)-b-
D-glucans, a major cell wall component together with
chitin and mannoproteins. Echinocandins inhibit the cat-
alytic subunit (Fksp) encoded by two or three FKS genes,
depending on the fungal species (Fig. 1)[7].
Azoles
Triazoles—fluconazole, itraconazole, voriconazole and
posaconazole—are the azoles most commonly used to
treat invasive fungal infections. Isavuconazole (ISA;
BAL4815), is a novel triazole currently in global phase 3
clinical trials for treatment of invasive fungal infections.
It showed good activity against Candida spp. with
reduced susceptibility to currently used azoles (personal
communications: Smart JI, P983, ECCMID, Berlin, 2013
and Maertens J., O230, ECCMID, Barcelona, 2014).
Triazoles block the synthesis of the main sterol of fungal
membranes, ergosterol, by targeting the lanosterol-14a-
demethylase, also called Erg11p or Cyp51p (Fig. 1). This
blockade has three major effects: (a) ergosterol depletion
and changes in membrane permeability, (b) changes in the
activity of membrane-bound proteins, some of which are
involved in cell wall synthesis and (c) synthesis of toxic
sterols as a result of Erg3p activity and accumulation of
14a-methylated sterols (Fig. 1)[8]. Azoles have long
been considered to act solely on the cell membrane, but
there is growing evidence to suggest that they also act on
the cell wall structure. Studies have demonstrated com-
pensatory responses similar to those observed with cell
wall-disrupting agents [9, 10].
Polyenes
Amphotericin B (AMB) and its lipid and liposomal
derivatives bind ergosterol, causing pore formation and
ion leakage, with fungicidal effects (Fig. 1). It has been
suggested that pore formation is not required for the
fungicidal effect, which is dependent only on ergosterol
binding [11]. In addition, a recent study revealed that
AMB is able to aggregate and to act like a ‘‘sponge’’, thus
extracting this key component from cell membranes [12].
Cholesterol is the major sterol of the mammalian mem-
branes. Ergosterol and cholesterol have different
structures, but drug specificity is not absolute and AMB
has also been shown to bind cholesterol [13]. New for-
mulations involving liposome encapsulation (L-AMB),
AMB colloidal dispersion (ABCD) and AMB lipid com-
plex (ABLC) have increased drug specificity and
delivery, greatly reducing toxicity without decreasing
efficacy [14]. However, ABCD caused a similar number
of infusion-related reactions to AMB, and is no longer
available.
Antifungal agents acting on nucleic acids and protein
synthesis
Flucytosine is a pyrimidine analogue that is converted
to 5-fluorouracil, which inhibits both RNA and DNA
synthesis. Cytosine permease (Fcy2p), cytosine deami-
nase (Fcy1p), and uracil phosphoribosyl transferase
(Fur1p) activities are required for antifungal activity
(Fig. 1).
Antifungal drug resistance in Candida spp.
Tolerance and resistance due to cellular stress
responses
An increase in cell wall chitin content has been shown to
occur in response to the exposure to echinocandin and
azoles in C. albicans [10, 15]. The blockade of this cell
wall compensatory mechanism with calcineurin or protein
kinase C (PKC) inhibitors restores the fungicidal activity
of both azoles and echinocandins consistent with the
hypothesis that chitin accumulation plays a role in toler-
ance to these drugs [16, 17]. Furthermore, a high chitin
content has been associated with resistance to echino-
candins in mice and in ‘paradoxical growth’, defined as
the ability to develop in vitro at high, but not intermediate
concentrations of a drug [18]. The clinical impact of this
paradoxical growth in vitro remains unclear, as it is also
related to lower virulence [19].
Molecular mechanisms of antifungal drug resistance
in Candida spp.
The molecular mechanisms of antifungal drug resistance
are presented in Fig. 2 and Table S1 (electronic supple-
mentary material).
Echinocandins
Molecular resistance to echinocandins is mediated
principally by mutations in FKS genes: FKS1 in Can-
dida spp., and FKS1 and FKS2 in C. glabrata. These
mutations are located in two‘‘hotspot’’regions,HS1
and HS2 and are mostly S645F/P/Y and S629P in FKS1
of C. albicans and C. glabrata respectively, and S663F/P
in C. glabrata FKS2 [7, 20]. These mutations confer
cross-resistance to all three echinocandins, by modify-
ing the catalytic and kinetic properties of the target
enzyme.
Azoles
Decreased susceptibility or resistance to azoles in
Candida spp. is mediated by various mechanisms,
which may operate simultaneously in a given isolate,
following sequential acquisition under drug pressure
[21, 22]. Drug efflux is a major mechanism, mediated
by mutations of genes encoding regulators of trans-
porters of the ATP-binding cassette (ABC) superfamily
or the major facilitator superfamily (MFS) [22, 23].
ABC transporter overexpression is associated with
cross-resistance to diverse azoles, whereas MFS trans-
porter overexpression is limited to resistance to fewer
Fig. 1 Targets and mechanisms of action of systemic antifungal
drugs. Sites and modes of action of the current classes of systemic
antifungal drugs used to treat invasive candidiasis. a Echinocandins
target cell wall synthesis, inhibiting (1,3)-b-
D-glucan synthesis,
which occurs on the inner side of the plasma membrane. b Azoles
target the ergosterol biosynthesis pathway in the endoplasmic
reticulum. They block 14a-demethylase (also called Erg11p or
Cyp51p), resulting in ergosterol depletion in the membrane and
activation of the Erg3p alternative pathway, leading to the synthesis
of toxic sterols. c Polyenes bind to cell membrane ergosterol
creating pores and aggregate, to act as a ‘‘sponge’’, thus resulting in
ion depletion. d Flucytosine acts in the nucleus, where its toxic
metabolites inhibit nucleic acid synthesis
azoles (fluconazole, voriconazole) (Table S1). A second
major mechanism is overproduction of the target
enzyme Erg11p [22]. Amino acid substitutions in
Erg11p may also decrease the affinity of the drugs for
this enzyme [24]. Finally, ERG3 mutations are associ-
ated with cross-resistance to azoles through a metabolic
bypass leading to the synthesis of fecosterol which is
able to replace ergosterol (Fig. 2)[25].
Other major genetic alterations may decrease azole
susceptibility. Aneuploidy, through chromosomal dupli-
cation or loss of heterozygosity, increases the copy
number of genes involved in azole resistance in C. albi-
cans and C. glabrata [22, 23, 26]. Respiratory and
mitochondrial deficiencies may also contribute to azole
resistance in these species [27].
Polyenes
Polyene resistance has been little described and the exact
mechanisms involved remain unclear, partly because of
the small number of clinical isolates displaying altered
susceptibility in vitro. Resistance is associated with
changes in membrane sterol composition due to mutations
in the genes of the ergosterol biosynthesis pathway:
ERG2, ERG3, ERG5, ERG6 and ERG11 [28].
Flucytosine
Two main mechanisms of flucytosine resistance are
known: (1) decreased uptake of the drug due to mutations
Fig. 2 Molecular mechanisms of echinocandin and azole resistance
in Candida spp. a Regular b-1,3-glucan synthesis on the inner side
of the fungal membrane. b Typical echinocandin activity. These
compounds block cell wall synthesis by inhibiting the Fksp subunit
of the b-1,3-glucan synthase. c Echinocandin resistance due to FKS
mutations. The target enzyme is less sensitive to echinocandins,
allowing the production of b-1,3-glucans. d Typical ergosterol
synthesis at the endosplamic reticulum and uptake of azole
antifungal drugs into the cytosol of the fungal cell. e Typical azole
activity. These molecules inhibit the lanosterol-14a-demethylase
(Erg11p), leading to (1) membrane ergosterol depletion and (2) the
production of toxic sterols via Erg3p. f Azole resistance due to (1)
the overproduction of transporters, increasing azole efflux, (2)
alteration of the target enzyme by mutations of ERG11, (3) Erg11p
overproduction, (4) mutations of ERG3 preventing the azole-
mediated production of toxic sterols which are substituted by the
non-toxic fecosterol
of the FCY2 gene encoding the cytosine permease, and (2)
impaired metabolism of the drug or its active metabolite
(5-FU) due to mutations of FCY1 or FUR1. Such muta-
tions have been described in clinical isolates of C.
albicans and C. lusitaniae [29, 30].
Antifungal drug resistance in Candida spp. biofilms
In ICUs, candidiasis may be favoured by biofilms for-
mation, mostly on catheters but also on other implanted
medical devices. Only a few antifungal drugs (L-AMB
and echinocandins) have some efficacy against yeasts
embedded in such complex structures [31]. Echinocandins
are active against biofilms, but are more effective against
biofilms containing C. albicans or C. glabrata than
against biofilms of C. tropicalis or C. parapsilosis [32,
33]. Conversely, yeast cells in biofilms are up to 1,000
times more resistant to azoles than their planktonic
counterparts [34].
The resistance of biofilms combines both planktonic
and biofilm-specific resistance mechanisms. Efflux pump
upregulation is involved in the early stages of biofilm
development, whereas the greater resistance of mature
biofilms is due to the presence of an extracellular matrix
(ECM) and persister cells, changes to the sterol compo-
sition of the membrane and the activation of stress-
induced pathways [35–37]. The ECM plays a key role, by
sequestering antifungal agents and preventing their
interaction with the target. This action is mediated at least
by (1,3)-b-
D-glucan polymers [38]. Stress responses,
mediated by the PKC, calcineurin and heat shock pro-
tein 90 (HSP90) pathways, also control ECM production
[35, 39]. Extracellular DNA also affects biofilm resistance
and the treatment of C. albicans biofilms with DNase
potentiates the antifungal activity of echinocandins and
polyenes, but not fluconazole [40]. Persister cells have
been described in Candida spp. biofilms, particularly
those formed by C. krusei and C. albicans. These cells are
phenotypic variants able to survive in the presence of
antifungal agents. They can again proliferate when drug
pressure is released and may cause relapses often
described in clinical situations [35, 37].
Laboratory detection of antifungal resistance
Antifungal drug susceptibility testing assays
Methods for in vitro susceptibility testing are available
from the Clinical and Laboratory Standards Institute
(CLSI) and the European Committee on Antimicrobial
Susceptibility Testing (EUCAST) [41, 42]. Other com-
mercially available standardised tests, such as Etest
"
(bioMe
´
rieux), Sensititre YeastOne
"
(TREK diagnostic
systems), ATB fungus 2
"
(bioMe
´
rieux) and Vitek-2
"
(bioMe
´
rieux), are more appropriate for routine clinical
use [43–45]. These tests determine the minimum inhibi-
tory concentration (MIC) or directly classify isolates as
susceptible (S), intermediate (I) or resistant (R), corre-
sponding to a high probability of treatment success (S), an
uncertain effect of treatment (I) or a high probability of
treatment failure (R). This classification is based on the
clinical breakpoints (CBPs) established for MIC inter-
pretation. Previous CBPs were not species-specific and
were too high to distinguish between C. glabrata isolates
susceptible and resistant to azoles and to detect emerging
resistance in C. albicans, C. tropicalis and C. parapsilo-
sis. In addition, clinical resistance to echinocandins due to
FKS mutations was increasingly being reported in patients
infected with ‘‘S’’ strains, defined with a former CBP
of 2 lg/ml or less. Up to 45 % of FKS mutants have been
incorrectly considered as susceptible [7, 46, 47].
The revised CBPs in current use are species-specific
and were established on the basis of five parameters: dose
regimens; MIC distributions from multiple laboratories;
epidemiologic cut-off values defined with respect to the
higher MIC of wild-type isolates; pharmacokinetic/phar-
macodynamic parameters and clinical outcome [20]. The
key changes concern the susceptibility of C. glabrata and
C. parapsilosis to fluconazole and echinocandins,
respectively. The ‘‘S’’ category was abolished by CLSI
and EUCAST for C. glabrata and fluconazole, consider-
ing all isolates to be intermediate or resistant. EUCAST
was even more severe in its approach, recommending, as
for C. krusei, that C. glabrata should not be tested with
fluconazole and that fluconazole should not be used for
C. glabrata infections [48]. The same removal of the ‘‘S’’
category was recommended, albeit only by EUCAST, for
C. parapsilosis and echinocandins [20]. One other major
difference between CLSI and EUCAST is that this latter
does not determine CBPs for caspofungin. Indeed, CLSI
and EUCAST agree that there is a lack of interlaboratory
reproducibility in MIC values for caspofungin. Until this
problem, not seen with other echinocandins, is resolved,
neither CLSI nor EUCAST recommends caspofungin
resistance testing [49–52]. EUCAST specifies that some
mutations decrease susceptibility to anidulafungin and
caspofungin but not micafungin, and thus recommends
the use of anidulafungin as a marker for echinocandin
resistance [20] (see Table 1 for simplified CLSI and
EUCAST updated CBPs). Both CLSI and EUCAST also
determined epidemiological cut-off values which are
more sensitive than CBP to detect non-wild-type isolates
exhibiting potential resistance mutations and
mechanisms.
AMB testing remains particularly challenging and
microbiological resistance is rarely detected [53]. Etest
"
(bioMe
´
rieux) was found to be superior to both CLSI and
EUCAST reference methods for identifying resistant and
intermediately susceptible C. glabrata isolates [54]. Thus,
AMB resistance is mainly identified through clinical
failure.
While current MIC testing protocols are adapted for
planktonic cells, these protocols are still not implemented
in biofilms. Since biofilms can be detected in infected
tissues, this is clearly another limitation in the interpre-
tation of susceptibility tests for predicting patient
outcome.
The performance of direct antifungal drug suscepti-
bility testing, through the use of Etest
"
(bioMe
´
rieux) on
positive blood samples, has been evaluated. Agreement
between direct and standard methods was high and false-
positive results for resistance to fluconazole and vorico-
nazole were obtained for 7 % of isolates, with false-
negative results obtained for 0.6 % of blood samples. No
errors were detected for caspofungin, but the method was
not reliable for AMB [55]. The new CBPs are species-
specific, so this approach requires a rapid identification
tool. Direct antifungal drug susceptibility testing should
therefore be re-evaluated, according to the current stan-
dards, for both categorisation and identification.
Molecular detection of mutations conferring
antifungal drug resistance
Molecular methods have been developed for the charac-
terisation of resistance-causing mutations. Culture-based
susceptibility assays take at least 24 h, but molecular
tools can assess resistance more rapidly and with greater
sensitivity. Both azole and echinocandin resistance
mutations are accurately detected with next-generation
sequencing platforms, allele-specific real-time probes,
melt curve analysis or microarrays, or microsphere-based
technologies, such as Luminex Mag Pix (Austin, TX)
[56–60] (C. Garnaud, personal communication). More-
over, as in Aspergillus fumigatus azole resistance, the
direct detection of mutations in clinical samples may
make it possible to detect mutations earlier by eliminating
the time required for culture [61].
Update on the epidemiology of Candida spp.
antifungal resistance
When focusing on species distribution and antifungal
resistance, recent epidemiological studies, including the
SENTRY cohort, did not show major differences between
ICU and non-ICU patients [62, 63]. In both ICU and non-
ICU, the five main species (i.e. C. albicans, C. parapsi-
losis, C. glabrata, C. tropicalis and C. krusei) are
responsible for more than 90 % of invasive fungal
infections [62–65]. C. albicans still stands in first place,
even if, since the early 2000s, a shift towards non-albi-
cans species was clearly noticed [66, 67]. The fluconazole
drug pressure may explain this trend but other factors,
mainly underlying conditions or antibacterial therapy,
have been suggested [68]. A main difference in species
distribution is related to geographical location. In
southern countries (Italy, Spain, South America)
C. parapsilosis ranks second while in northern countries
C. glabrata is the most frequent species after C. albicans
[69]. These site specificities highlight the importance of
local data on Candida epidemiology, specific to each
health-care centre.
Table 1 EUCAST and CLSI antifungal breakpoints for the main Candida species
Antifungal agent MIC breakpoint (mg/l)
C. albicans C. glabrata C. krusei C. parapsilosis C. tropicalis
BS [R BS [R BS [R BS [R BS [R
Amphotericin B
EUCAST 1 1 1 1 1 1 1 1 1 1
CLSI ND ND ND ND ND ND ND ND ND ND
Fluconazole
EUCAST 2 4 0.002 32 – – 2 4 2 4
CLSI 2 4 0.002 32 – – 2 4 2 4
Voriconazole
EUCAST 0.12 0.12 IE IE IE IE 0.12 0.12 0.12 0.12
CLSI 0.12 0.5 – – 0.5 1 0.12 0.5 0.12 0.5
Anidulafungin
EUCAST 0.03 0.03 0.06 0.06 0.06 0.06 0.002 4 0.06 0.06
CLSI 0.25 0.5 0.12 0.25 0.25 0.5 2 4 0.25 0.5
Micafungin
EUCAST 0.016 0.016 0.03 0.03 IE IE 0.002 2 IE IE
CLSI 0.25 0.5 0.06 0.12 0.25 0.5 2 4 0.25 0.5
Adapted from Arendrup et al. [20] drug resistance updates (doi:10.1016/j.drup.2014.01.001) with permission. For complete data, see
Arendrup et al. [20]
ND not done, IE insufficient evidence
Indeed, some non-albicans species show intrinsic
resistance. For example C. glabrata and C. krusei are less
susceptible to azoles than other species (Table 2;[70–72])
and C. parapsilosis is less susceptible to echinocandins
owing to naturally occurring polymorphisms of the FKS
genes [73, 74]. Breakthrough infections with these species
may therefore occur during azoles or echinocandin
treatment [75, 76].
Another major and increasing threat is the risk of
becoming infected with a strain which has acquired a
resistant phenotype. Acquired resistance is thought to be
rare in Candida spp., or at least less frequent than intrinsic
resistance. Fortunately, yeasts, unlike bacteria, do not
display the horizontal transmission of resistance genes
[77]. Moreover, cross-contamination between patients
and health-care workers has been described mostly for C.
albicans and C. parapsilosis but remains rare [69, 78, 79].
Acquired resistance thus results principally from the
selection of mutants subjected to drug pressure in
patients.
Acquired resistance to echinocandins is increasingly
reported for most of the clinically important Candida
spp. It remains uncommon in C. albicans (\1 %), C.
tropicalis (\5 %) and C. krusei (\7 %), but is now
becoming frequent in C. glabrata (8–15 %) [5, 63, 80,
81]. One recent study showed that the frequency of
echinocandin resistance in C. glabrata increased from
4.9 to 12.3 % between 2001 and 2010 [5]. It has been
shown that 7 days of exposure to echinocandin is suf-
ficient to induce FKS mutations in C. glabrata [5, 6].
The haploid trait of this species may partly explain the
higher level of expression of molecular resistance
exhibited by C. glabrata. FKS mutations have been
described in almost all the clinically important Candida
species:
C. albicans, C. glabrata, C. tropicalis [82, 83],
C. krusei [84] and C. kefyr [85] and breakthrough
infections are also increasingly reported [7, 75, 76, 86–
90]. A recent study focusing on C. glabrata candidemia
described 18 % of FKS mutation, with prior echino-
candin exposure as the only independent risk factor for
the development of these mutations [91] confirming the
results obtained previously by Alexander et al. [5].
Interestingly, the nature and/or the number of FKS
mutations in C. glabrata and C. albicans influences the
level of resistance in vivo [91, 92]. Even if the micro-
biological resistance to echinocandins is still uncommon,
the growing incidence of FKS mutations is worrying and
needs to be very closely monitored. FKS resistance
mutations also need to be more deeply studied.
Azoles and especially fluconazole are widely pre-
scribed for ICU patients. Acquired fluconazole resistance
is frequent in C. glabrata (from 4 to 16 %), which
increasingly displays cross-resistance to voriconazole. So
far, multidrug-resistant phenotype against azole and ech-
inocandins has only been described for C. glabrata and is
a matter of serious concern [5, 63, 66, 80, 81, 93, 94].
Fluconazole resistance remains uncommon in C. albicans
(\5 %), but is more prevalent in C. parapsilosis (4–10 %)
and C. tropicalis (4–9 %) [63, 64, 81]. However, the
recent China-SCAN study reported higher rates of fluco-
nazole resistance in C. albicans (9.6 %) and C.
parapsilosis (19.3 %) which may reflect geographical
differences [93]. Again, most studies report that a previ-
ous history of azole pre-exposure increases the risk of
in vitro azole resistance (from 2 to 58 % in a 2013 study
by Montagna et al. [65]).
Resistance to AMB remains rare despite its use in
monotherapy for years. This may be due to its inherently
fungicidal effect, limiting the selection of mutants.
However, resistant isolates of C. glabrata and C. krusei
Table 2 Spectrum of activity of the antifungal agents used to treat invasive candidiasis
Candida spp. Polyenes Azoles Echinocandins Flucytosine
AMB formulations FLU ITRA VOR POSA CAS MIC ANI
C. albicans ?? ?? ?? ?? ?? ?? ?? ?? ??
C. glabrata ?
a
?/-?/- ? ? ?? ?? ?? ??
C. parapsilosis ?? ?? ?? ?? ?? ? ? ? ??
C. tropicalis ?? ?? ?? ?? ?? ?? ?? ?? ??
C. krusei ? 2 ?/- ? ? ?? ?? ?? 2
C. rugosa ?
a
? ? ?? ?? ? ? ? ??
C. guilliermondii ?? ?? ?? ?? ?? ? ? ? ??
C. lusitaniae ?? ?? ?? ?? ?? ?? ?? ?? ??
C. inconspicua ?? - ?/- ? ? ?? ?? ?? NS
C. norvegensis ?? - ?/-?/-?/- ?? ?? ?? NS
Adapted with permission from Denning DW, Hope WW (2010)
Trends Microbiol (doi:10.1016/j.tim.2010.02.004). In vitro inherent
activity: ?? good activity, ? mild activity, ?/- slight activity,
- no activity; NS not specified
AMB amphotericin B, FLU fluconazole, ITRA itraconazole, VOR
voriconazole, POSA posaconazole, CAS caspofungin, MIC mica-
fungin, ANI anidulafungin
a
This slight decrease in susceptibility to AMB is more pronounced
in North America than in Europe
are increasingly being reported and this new entity also
needs to be closely monitored [54, 95–98].
Even if it remains uncommon, Candida spp. drug
resistance is clearly becoming an ‘every-day’ concern in
the mycology laboratory. Determining initial but also
subsequent MICs is necessary to assess microbial resis-
tance emergence.
Clinical impact of antifungal resistance
Clinical resistance
The failure of antifungal therapy or clinical resistance is
defined as a steady-to-worse infectious syndrome with no
improvement of attributable symptoms during the evalu-
ation, death being the ‘‘ultimate’’ failure. However, it
remains difficult to assess whether the patient dies with or
of fungal infection. These criteria classify clinical out-
come in trials, but can also be applied for bedside
management [99]. Breakthrough infections are considered
as clinical resistance and are microbiologically docu-
mented. They have frequently, but not exclusively, been
described in cases of echinocandin exposure [5, 7, 75, 76,
86–90, 100, 101].
Despite its recent spread, microbiological resistance is
not the major factor underlying clinical resistance.
Indeed, underlying diseases, immunosuppression, com-
plicated abdominal surgery, extreme age and renal failure,
all frequently encountered in ICU patients, are known to
be predictors of mortality in cases of invasive candidiasis
[102, 103]. Clinical failure may also occur when the
effective concentration of the chosen drug is not reached
at the infected site. This situation frequently occurs for
biofilms on prosthetic devices or catheters, abscesses,
chorioretinitis or endophthalmitis, or other sanctuary foci.
As a result of their multiple comorbidities and manage-
ment strategies, ICU patients may display higher
pharmacokinetic (PK) instability than other patients.
Thus, regular, complete investigations of deep infections
and assessments of the PK/pharmacodynamic properties
of antifungal drugs are essential for correct appraisal of
the clinical response. For example, given their poor
penetration into the eye, echinocandins are not recom-
mended in cases of suspected ocular secondary
dissemination of Candida, whereas echinocandins or
L-AMB are the drugs of choice when central catheters
cannot be removed [104]. Given the lower frequency of
primary resistance than initially thought and its beneficial
penetration properties, 5-fluorocytosine may be adminis-
tered in combination with other drugs to treat invasive
candidiasis at deep secondary sites [105]. A recent review
has provided a comprehensive analysis of the tissue
penetration properties of current antifungal agents [106].
Clinical relevance of in vitro antifungal drug
susceptibility testing and of the molecular detection
of mutations
Crude MICs are not sufficient to predict clinical outcome.
Candidemia due to C. tropicalis, for which the MICs of all
antifungal agents are very low, has been associated with a
higher mortality than for other species [102]. By contrast,
C. parapsilosis isolates have high MICs for echinocandins,
although treatment failure remains rare [104, 107]. These
discrepancies between laboratory tests on antifungal drugs
and clinical outcome have been extensively reported and
are due to several factors, including a species-dependent
virulence traits and patient-dependent conditions. The use
of the revised CBPs, partly taking these factors into
account, may improve the clinical predictive value of
in vitro susceptibility tests. Thus, close monitoring of MICs
together with accurate interpretation based on revised
CBPs is always warranted to ensure appropriate specific
treatment (Fig. 3). Antifungal susceptibility testing (AST)
on Candida strains isolated from deep sites is recom-
mended by the European Society of Clinical Microbiology
and Infectious Diseases (ESCMID) [104]. Reference
methods are preferred but commercial techniques can be
used if verification has been made that the endpoint for each
species mirrors those of reference methods [104].
There is growing evidence that the detection of muta-
tions, and especially FKS mutations, could be used as a
predictive marker of clinical failure. In one recent study,
FKS mutations were found in 7.9 % of 313 C. glabrata
isolates from blood samples, and up to 80 % of patients
infected with strains with both FKS mutations and high
MICs for caspofungin experienced clinical failure or
recurrent infection [5]. Another study identified C. glab-
rata FKS mutation as the only independent risk factor
associated with clinical failure and showed that the
detection of FKS mutations was superior to MIC for pre-
dicting treatment response [108]. The same group
subsequently showed that the Etest
"
method (bioMe
´
rieux,
France) and a MIC greater than 0.25 ll/ml for caspofungin
provided 100 % sensitivity and 94 % specificity for the
identification of FKS mutant isolates. Prior echinocandin
exposure and MIC values greater than 0.25, 0.06 and
0.03 ll/ml for caspofungin, anidulafungin and micafun-
gin, respectively, were found to be predictive of clinical
failure in 91, 89 and 78 % of patients with treatment
failure, respectively [6]. These findings have led to valu-
able, easy-to-use algorithms for predicting the outcome of
echinocandin treatment from MIC levels and prior echi-
nocandin exposure status. Not all mycology laboratories
are equipped to detect FKS mutations and the Etest
"
method performs well for the detection of non-wild-type
strains [109, 110]; this bedside strategy can therefore be
used to identify patients at risk of treatment failure, for
whom other antifungal treatments should be prescribed.
Similar predictive markers have been suggested for
azole resistance. Exposure to fluconazole in the last
30 days has been shown to have a significant impact on
species distribution and MIC [4]. Algorithms have also
been developed for assessment of the growing risk of C.
glabrata infections. Cohen et al. identified six indepen-
dent risk factors for C. glabrata fungemia in ICU patients:
age greater than 60 years, recent abdominal surgery, less
than 7 days between ICU admission and first positive
blood culture, recent use of cephalosporins, solid tumour
and absence of diabetes mellitus [111].
Integration of clinical and microbiological data (as
proposed in Fig. 3) is thus crucial to improve the pre-
diction of treatment response. Previous exposure to
fluconazole and echinocandin should be accurately mon-
itored, although the exact period to be considered remains
to be defined. Patients receiving prophylactic, empirical
or targeted antifungal therapy should be carefully moni-
tored for breakthrough infections. Local epidemiological
investigations and MIC determinations for Candida spp.
isolates are also crucial and should be interpreted with the
most appropriate, revised CBPs. Molecular tools are also
required for the rapid detection of mutant strains.
Impact of antifungal drug resistance on patient
management
Epidemiological changes have a direct impact on clinical
management, leading to the updating of international
expert committee recommendations [104, 112–116].
These recommendations propose consensual attitudes to
the management of invasive candidiasis, but divergence
remains on several crucial, contentious points [107, 117,
118], which may be confusing for clinicians treating
patients.
All experts agree that patients with Candida-positive
blood cultures should be treated with systemic antifungal
drugs, but ESCMID cites echinocandins as the only initial
treatment with the highest levels of strength of recom-
mendation (A) and quality of evidence (I) [104], whereas
the European Conference on Infections in Leukaemia
(ECIL) and the Infectious Diseases Society of America
(IDSA) consider fluconazole at the AI level of recom-
mendation as a suitable alternative for patients with less
severe or stable infection not previously exposed to azoles
[10, 115, 116]. Indeed, in a recent study including 216
patients with Candida-induced septic shock, no difference
Fig. 3 Bedside strategy for circumventing antifungal drug resis-
tance in 2014. ATF antifungal drug, FCZ fluconazole, CAS
caspofungin, PK/PD pharmacokinetics and pharmacodynamics,
TDM therapeutic drug monitoring, MIC minimal inhibitory
concentration, CBP clinical breakpoint
in mortality was observed between patients treated with
fluconazole or with echinocandins [119]. In patients at
risk of C. glabrata candidemia, echinocandins should be
preferred. Voriconazole is not usually used as first-line
therapy but it offers an alternate option for intrinsically
less susceptible species (C. krusei or C. glabrata).
Because acquired mutations can lead to cross-resistance
to both fluconazole and voriconazole, a strain resistant to
fluconazole should not be treated with voriconazole
unless its susceptibility profile has been confirmed, or the
mutation genetically characterized.
All expert panels strongly recommend the removal of
central venous catheters ‘whenever possible’, but ESC-
MID guidelines suggest that replacement is not formally
required in patients treated with echinocandins or L-AMB
[104]. Catheter exchange via a guide wire entails a risk of
contaminating the new device with Candida and should
be restricted to patients with limited venous access [107,
116, 120]. Given the specific link between C. parapsilosis
and catheter infections and the low susceptibility of this
species to echinocandins, catheter removal is appropriate
in patients with invasive C. parapsilosis candidiasis. In
stabilised patients infected with a fluconazole-susceptible
isolate, with negative blood cultures, step-down therapy
onto oral fluconazole is recommended, over a period of
3–10 days, depending on the guidelines considered.
Conclusions
Although drug resistance is rapidly spreading in Candida
spp., antifungal treatments are still generally successful:
up to 80 % of C. albicans infections are cleared with
echinocandins. Treatment success rates are also generally
satisfactory for fluconazole. However, the emergence of
antifungal resistance must be considered at the patient
level in order to improve patient management. In ICUs,
intrinsic resistance of C. glabrata and C. krusei to
fluconazole can be detected and handled rapidly through
correct species identification, detailed assessment of
antifungal drug exposure and Candida spp. colonisation
history. The emergence of acquired resistance during or
after treatment is more worrying: it mostly involves C.
glabrata and the echinocandins and leads to breakthrough
infections or treatment failures. This highlights the need
for (a) accurate assessments of prior antifungal exposure,
(b) close monitoring of patients on antifungal drugs,
(c) the routine surveillance of in vitro susceptibility test-
ing and (d) the development of feasible methods for rapid
detection of mutations. The emergence of resistance
should also be considered at the community level as a
warning sign triggering improvements in antifungal drug
use, particularly in patients for whom the potential benefit
of treatment is unproven. Closer monitoring of antifungal
drug use is thus required.
Acknowledgments We are grateful to Audrey Le Goue
¨
llec for her
assistance in preparing the figures.
Conflicts of interest D. Maubon, C. Garnaud and M. Cornet
received a research grant from Pfizer in 2013. T. Calandra: board
membership: Pfizer; Consultancy: Pfizer, MSD; Speakers bureaus:
BioMe
´
rieux, Pfizer; Development & educational presentations:
MSD, Gilead Sciences (money to institution); Travel & meeting
expenses: Astellas, Pfizer.
References
1. Leo
´
n C, Ostrosky-Zeichner L,
Schuster M (2014) What’s new in the
clinical and diagnostic management of
invasive candidiasis in critically ill
patients. Intensive Care Med. doi:
10.1007/s00134-014-3281-0
2. Guery BP, Arendrup MC, Auzinger G
et al (2009) Management of invasive
candidiasis and candidemia in adult
non-neutropenic intensive care unit
patients: part II. Treatment Intensive
Care Med 35:206–214. doi:
10.1007/s00134-008-1339-6
3. Fournier P, Schwebel C, Maubon D
et al (2011) Antifungal use influences
Candida species distribution and
susceptibility in the intensive care
unit. J Antimicrob Chemother
66:2880–2886
4. Lortholary O, Desnos-Ollivier M,
Sitbon K et al (2011) Recent exposure
to caspofungin or fluconazole
influences the epidemiology of
candidemia: a prospective multicenter
study involving 2,441 patients.
Antimicrob Agents Chemother
55:532–538. doi:
10.1128/AAC.01128-10
5. Alexander BD, Johnson MD, Pfeiffer
CD et al (2013) Increasing
echinocandin resistance in Candida
glabrata: clinical failure correlates
with presence of FKS mutations and
elevated minimum inhibitory
concentrations. Clin Infect Dis
56:1724–1732. doi:10.1093/cid/cit136
6. Shields RK, Nguyen MH, Press EG
et al (2013) Caspofungin MICs
correlate with treatment outcomes
among patients with Candida glabrata
invasive candidiasis and prior
echinocandin exposure. Antimicrob
Agents Chemother 57:3528–3535. doi:
10.1128/AAC.00136-13
7. Perlin DS (2011) Current perspectives
on echinocandin class drugs. Future
Microbiol 6:441–457. doi:
10.2217/fmb.11.19
8. Odds FC, Brown AJP, Gow NAR
(2003) Antifungal agents: mechanisms
of action. Trends Microbiol
11:272–279
9. Sorgo AG, Heilmann CJ, Dekker HL
et al (2011) Effects of fluconazole on
the secretome, the wall proteome, and
wall integrity of the clinical fungus
Candida albicans. Eukaryot Cell
10:1071–1081. doi:
10.1128/EC.05011-11
10. Pfaller M, Riley J (1992) Effects of
fluconazole on the sterol and
carbohydrate composition of four
species of Candida. Eur J Clin
Microbiol Infect Dis 11:152–156
11. Gray KC, Palacios DS, Dailey I et al
(2012) Amphotericin primarily kills
yeast by simply binding ergosterol.
Proc Natl Acad Sci U S A
109:2234–2239. doi:
10.1073/pnas.1117280109
12. Anderson TM, Clay MC, Cioffi AG
et al (2014) Amphotericin forms an
extramembranous and fungicidal
sterol sponge. Nat Chem Biol
10:400–406. doi:
10.1038/nchembio.1496
13. Kotler-Brajtburg J, Price HD, Medoff
G et al (1974) Molecular basis for the
selective toxicity of amphotericin B
for yeast and filipin for animal cells.
Antimicrob Agents Chemother
5:377–382
14. Hamill RJ (2013) Amphotericin B
formulations: a comparative review of
efficacy and toxicity. Drugs
73:919–934. doi:
10.1007/s40265-013-0069-4
15. Walker LA, Gow NAR, Munro CA
(2010) Fungal echinocandin
resistance. Fungal Genet Biol
47:117–126. doi:
10.1016/j.fgb.2009.09.003
16. Sanglard D, Ischer F, Marchetti O et al
(2003) Calcineurin A of Candida
albicans: involvement in antifungal
tolerance, cell morphogenesis and
virulence. Mol Microbiol 48:959–976
17. LaFayette SL, Collins C, Zaas AK
et al (2010) PKC signaling regulates
drug resistance of the fungal pathogen
Candida albicans via circuitry
comprised of Mkc1, calcineurin, and
Hsp90. PLoS Pathog 6:e1001069. doi:
10.1371/journal.ppat.1001069
18. Lee KK, Maccallum DM, Jacobsen
MD et al (2012) Elevated cell wall
chitin in Candida albicans confers
echinocandin resistance in vivo.
Antimicrob Agents Chemother
56:208–217. doi:
10.1128/AAC.00683-11
19. Rueda C, Cuenca-Estrella M,
Zaragoza O (2014) Paradoxical
growth of Candida albicans in the
presence of caspofungin is associated
with multiple cell wall rearrangements
and decreased virulence. Antimicrob
Agents Chemother 58:1071–1083. doi:
10.1128/AAC.00946-13
20. Arendrup MC, Cuenca-Estrella M,
Lass-Flo
¨
rl C, Hope WW (2014)
Breakpoints for antifungal agents: an
update from EUCAST focussing on
echinocandins against Candida spp
and triazoles against Aspergillus spp.
Drug Resist Updat 16(6):81–95. doi:
10.1016/j.drup.2014.01.001
21. Coste A, Selmecki A, Forche A et al
(2007) Genotypic evolution of azole
resistance mechanisms in sequential
Candida albicans isolates. Eukaryot
Cell 6:1889–1904
22. Sanglard D, Coste A, Ferrari S (2009)
Antifungal drug resistance
mechanisms in fungal pathogens from
the perspective of transcriptional gene
regulation. FEMS Yeast Res
9:1029–1050. doi:
10.1111/j.1567-1364.2009.00578.x
23. Coste A, Turner V, Ischer F et al
(2006) A mutation in Tac1p, a
transcription factor regulating CDR1
and CDR2, is coupled with loss of
heterozygosity at chromosome 5 to
mediate antifungal resistance in
Candida albicans. Genetics
172:2139–2156. doi:
10.1534/genetics.105.054767
24. Morio F, Loge C, Besse B et al (2010)
Screening for amino acid substitutions
in the Candida albicans Erg11 protein
of azole-susceptible and azole-
resistant clinical isolates: new
substitutions and a review of the
literature. Diagn Microbiol Infect Dis
66:373–384. doi:
10.1016/j.diagmicrobio.2009.11.006
25. Morio F, Pagniez F, Lacroix C et al
(2012) Amino acid substitutions in the
Candida albicans sterol D5,6-
desaturase (Erg3p) confer azole
resistance: characterization of two
novel mutants with impaired
virulence. J Antimicrob Chemother
67:2131–2138. doi:
10.1093/jac/dks186
26. Pola
´
kova
´
S, Blume C, Za
´
rate JA et al
(2009) Formation of new
chromosomes as a virulence
mechanism in yeast Candida glabrata.
Proc Natl Acad Sci U S A
106:2688–2693. doi:
10.1073/pnas.0809793106
27. Ferrari S, Sanguinetti M, De Bernardis
F et al (2011) Loss of mitochondrial
functions associated with azole
resistance in Candida glabrata results
in enhanced virulence in mice.
Antimicrob Agents Chemother
55:1852–1860. doi:
10.1128/AAC.01271-10
28. Vincent BM, Lancaster AK, Scherz-
Shouval R et al (2013) Fitness trade-
offs restrict the evolution of resistance
to amphotericin B. PLoS Biol
11:e1001692. doi:
10.1371/journal.pbio.1001692
29. Spampinato C, Leonardi D (2013)
Candida infections, causes, targets,
and resistance mechanisms: traditional
and alternative antifungal agents. Bio
Med Res Int 2013:204237. doi:
10.1155/2013/204237
30. Florent M, Noe
¨
l T, Ruprich-Robert G
et al (2009) Nonsense and missense
mutations in FCY2 and FCY1 genes
are responsible for flucytosine
resistance and flucytosine-fluconazole
cross-resistance in clinical isolates of
Candida lusitaniae. Antimicrob
Agents Chemother 53:2982–2990. doi:
10.1128/AAC.00880-08
31. Delattin N, Cammue BPA, Thevissen
K (2014) Reactive oxygen species-
inducing antifungal agents and their
activity against fungal biofilms. Future
Med Chem 6:77–90. doi:
10.4155/fmc.13.189
32. Kuhn DM, George T, Chandra J et al
(2002) Antifungal susceptibility of
Candida biofilms: unique efficacy of
amphotericin B lipid formulations and
echinocandins. Antimicrob Agents
Chemother 46:1773–1780
33. Choi HW, Shin JH, Jung SI et al
(2007) Species-specific differences in
the susceptibilities of biofilms formed
by Candida bloodstream isolates to
echinocandin antifungals. Antimicrob
Agents Chemother 51:1520–1523. doi:
10.1128/AAC.01141-06
34. Lamfon H, Porter SR, McCullough M,
Pratten J (2004) Susceptibility of
Candida albicans biofilms grown in a
constant depth film fermentor to
chlorhexidine, fluconazole and
miconazole: a longitudinal study.
J Antimicrob Chemother 53:383–385.
doi:10.1093/jac/dkh071
35. Taff HT, Mitchell KF, Edward JA,
Andes DR (2013) Mechanisms of
Candida biofilm drug resistance.
Future Microbiol 8:1325–1337. doi:
10.2217/fmb.13.101
36. Tobudic S, Kratzer C, Lassnigg A,
Presterl E (2012) Antifungal
susceptibility of Candida albicans in
biofilms. Mycoses 55:199–204. doi:
10.1111/j.1439-0507.2011.02076.x
37. Mathe
´
L, Van Dijck P (2013) Recent
insights into Candida albicans biofilm
resistance mechanisms. Curr Genet.
doi:10.1007/s00294-013-0400-3
38. Mitchell KF, Taff HT, Cuevas MA
et al (2013) Role of matrix b-1,3
glucan in antifungal resistance of non-
albicans Candida biofilms.
Antimicrob Agents Chemother
57:1918–1920. doi:
10.1128/AAC.02378-12
39. Robbins N, Uppuluri P, Nett J et al
(2011) Hsp90 governs dispersion and
drug resistance of fungal biofilms.
PLoS Pathog 7:e1002257. doi:
10.1371/journal.ppat.1002257
40. Martins M, Henriques M, Lopez-Ribot
JL, Oliveira R (2012) Addition of
DNase improves the in vitro activity of
antifungal drugs against Candida
albicans biofilms. Mycoses 55:80–85.
doi:
10.1111/j.1439-0507.2011.02047.x
41. EUCAST (2008) EUCAST definitive
document EDef 7.1: method for the
determination of broth dilution MICs
of antifungal agents for fermentative
yeasts. Clin Microbiol Infect
14:398–405
42. CLSI Clinical and Laboratory
Standards Institute (2008) Reference
method for broth dilution antifungal
susceptibility testing of yeasts. Third
informational supplement. CLSI
document M27-S3. Clinical and
Laboratory Standards Institute, Wayne
43. Chryssanthou E, Cuenca-Estrella M
(2002) Comparison of the antifungal
susceptibility testing subcommittee
of the European Committee on
Antibiotic Susceptibility Testing
proposed standard and the E-test with
the NCCLS broth microdilution
method for voriconazole and
caspofungin susceptibility testing of
yeast species. J Clin Microbiol
40:3841–3844
44. Cuenca-Estrella M, Gomez-Lopez A,
Alastruey-Izquierdo A et al (2010)
Comparison of the Vitek 2 antifungal
susceptibility system with the clinical
and laboratory standards institute
(CLSI) and European Committee on
Antimicrobial Susceptibility Testing
(EUCAST) broth microdilution
reference methods and with the
sensititre yeastone and Etest
techniques for in vitro detection of
antifungal resistance in yeast isolates.
J Clin Microbiol 48:1782–1786. doi:
10.1128/JCM.02316-09
45. Lombardi G, Farina C, Andreoni S
et al (2004) Comparative evaluation of
Sensititre YeastOne vs. the NCCLS
M27A protocol and E-test for
antifungal susceptibility testing of
yeasts. Mycoses 47:397–401. doi:
10.1111/j.1439-0507.2004.01013.x
46. Pfaller MA, Diekema DJ, Andes D
et al (2011) Clinical breakpoints for
the echinocandins and Candida
revisited: integration of molecular,
clinical, and microbiological data to
arrive at species-specific interpretive
criteria. Drug Resist Updat
14:164–176. doi:
10.1016/j.drup.2011.01.004
47. Arendrup MC, Garcia-Effron G,
Buzina W et al (2009) Breakthrough
Aspergillus fumigatus and Candida
albicans double infection during
caspofungin treatment: laboratory
characteristics and implication for
susceptibility testing. Antimicrob
Agents Chemother 53:1185–1193. doi:
10.1128/AAC.01292-08
48. Pfaller MA, Andes D, Diekema DJ
et al (2010) Wild-type MIC
distributions, epidemiological cutoff
values and species-specific clinical
breakpoints for fluconazole and
Candida: time for harmonization of
CLSI and EUCAST broth
microdilution methods. Drug Resist
Updat 13:180–195. doi:
10.1016/j.drup.2010.09.002
49. Espinel-Ingroff A, Arendrup MC,
Pfaller MA et al (2013) Interlaboratory
variability of caspofungin MICs for
Candida spp. Using CLSI and
EUCAST methods: should the clinical
laboratory be testing this agent?
Antimicrob Agents Chemother
57:5836–5842. doi:
10.1128/AAC.01519-13
50. Pfaller MA, Messer SA, Diekema DJ
et al (2014) Use of micafungin as a
surrogate marker to predict
susceptibility and resistance to
caspofungin among 3,764 clinical
isolates of Candida by use of CLSI
methods and interpretive criteria.
J Clin Microbiol 52:108–114. doi:
10.1128/JCM.02481-13
51. Eschenauer GA, Nguyen MH, Shoham
S et al (2014) Real-world experience
with echinocandin MICs against
Candida species in a multicenter study
of hospitals that routinely perform
susceptibility testing of bloodstream
isolates. Antimicrob Agents
Chemother 58:1897–1906. doi:
10.1128/AAC.02163-13
52. Shields RK, Nguyen MH, Press EG
et al (2013) Anidulafungin and
micafungin minimum inhibitory
concentration breakpoints are superior
to caspofungin for identifying FKS
mutant Candida glabrata and
echinocandin resistance. Antimicrob
Agents Chemother. doi:
10.1128/AAC.01451-13
53. Park BJ, Arthington-Skaggs BA,
Hajjeh RA et al (2006) Evaluation of
amphotericin B interpretive
breakpoints for Candida bloodstream
isolates by correlation with therapeutic
outcome. Antimicrob Agents
Chemother 50:1287–1292. doi:
10.1128/AAC.50.4.1287-1292.2006
54. Krogh-Madsen M, Arendrup MC,
Heslet L, Knudsen JD (2006)
Amphotericin B and caspofungin
resistance in Candida glabrata isolates
recovered from a critically ill patient.
Clin Infect Dis 42:938–944. doi:
10.1086/500939
55. Guinea J, Recio S, Escribano P et al
(2010) Rapid antifungal susceptibility
determination for yeast isolates by use
of Etest performed directly on blood
samples from patients with fungemia.
J Clin Microbiol 48:2205–2212. doi:
10.1128/JCM.02321-09
56. Wiederhold NP, Grabinski JL, Garcia-
Effron G et al (2008) Pyrosequencing
to detect mutations in FKS1 that
confer reduced echinocandin
susceptibility in Candida albicans.
Antimicrob Agents Chemother
52:4145–4148. doi:
10.1128/AAC.00959-08
57. Park S, Perlin DS (2005) Establishing
surrogate markers for fluconazole
resistance in Candida albicans.
Microb Drug Resist 11:232–238. doi:
10.1089/mdr.2005.11.232
58. Loeffler J, Hagmeyer L, Hebart H et al
(2000) Rapid detection of point
mutations by fluorescence resonance
energy transfer and probe melting
curves in Candida species. Clin Chem
46:631–635
59. Wang H, Kong F, Sorrell TC et al
(2009) Rapid detection of ERG11
gene mutations in clinical Candida
albicans isolates with reduced
susceptibility to fluconazole by rolling
circle amplification and DNA
sequencing. BMC Microbiol 9:167.
doi:10.1186/1471-2180-9-167
60. Pham CD, Bolden CB, Kuykendall RJ,
Lockhart SR (2013) Development of a
Luminex-based multiplex assay for
detection of mutations conferring
resistance to echinocandins in
Candida glabrata. J Clin Microbiol.
doi:10.1128/JCM.03378-13
61. Zhao Y, Stensvold CR, Perlin DS,
Arendrup MC (2013) Azole resistance
in Aspergillus fumigatus from
bronchoalveolar lavage fluid samples
of patients with chronic diseases.
J Antimicrob Chemother
68:1497–1504. doi:10.1093/jac/dkt071
62. Bassetti M, Merelli M, Righi E et al
(2013) Epidemiology, species
distribution, antifungal susceptibility,
and outcome of candidemia across five
sites in Italy and Spain. J Clin
Microbiol 51:4167–4172. doi:
10.1128/JCM.01998-13
63. Pfaller MA, Messer SA, Moet GJ et al
(2011) Candida bloodstream
infections: comparison of species
distribution and resistance to
echinocandin and azole antifungal
agents in intensive care unit (ICU) and
non-ICU settings in the SENTRY
antimicrobial surveillance program
(2008–2009). Int J Antimicrob Agents
38:65–69. doi:
10.1016/j.ijantimicag.2011.02.016
64. Puig-Asensio M, Pema
´
n J, Zaragoza R
et al (2014) Impact of therapeutic
strategies on the prognosis of
candidemia in the ICU. Crit Care Med
42:1423–1432. doi:
10.1097/CCM.0000000000000221
65. Montagna MT, Caggiano G, Lovero G
et al (2013) Epidemiology of invasive
fungal infections in the intensive care
unit: results of a multicenter Italian
survey (AURORA Project). Infection
41:645–653. doi:
10.1007/s15010-013-0432-0
66. Diekema D, Arbefeville S, Boyken L
et al (2012) The changing
epidemiology of healthcare-associated
candidemia over three decades. Diagn
Microbiol Infect Dis 73:45–48. doi:
10.1016/j.diagmicrobio.2012.02.001
67. Chow JK, Golan Y, Ruthazer R et al
(2008) Factors associated with
candidemia caused by non-albicans
Candida species versus Candida
albicans in the intensive care unit.
Clin Infect Dis 46:1206–1213. doi:
10.1086/529435
68. Kanafani ZA, Perfect JR (2008)
Resistance to antifungal agents:
mechanisms and clinical impact. Clin
Infect Dis 46:120–128. doi:
10.1086/524071
69. Guinea J (2014) Global trends in the
distribution of Candida species
causing candidemia. Clin Microbiol
Infect. doi:10.1111/1469-0691.12539
70. Pfaller MA, Diekema DJ (2010)
Epidemiology of invasive mycoses in
North America. Crit Rev Microbiol
36:1–53
71. Guitard J, Angoulvant A, Letscher-
Bru V et al (2013) Invasive infections
due to Candida norvegensis and
Candida inconspicua: report of 12
cases and review of the literature. Med
Mycol. doi:
10.3109/13693786.2013.807444
72. Pfaller MA, Diekema DJ, Colombo
AL et al (2006) Candida rugosa, an
emerging fungal pathogen with
resistance to azoles: geographic and
temporal trends from the ARTEMIS
DISK antifungal surveillance program.
J Clin Microbiol 44:3578–3582. doi:
10.1128/JCM.00863-06
73. Garcia-Effron G, Katiyar SK, Park S
et al (2008) A naturally occurring
proline-to-alanine amino acid change
in FKS1p in Candida parapsilosis,
Candida orthopsilosis, and Candida
metapsilosis accounts for reduced
echinocandin susceptibility.
Antimicrob Agents Chemother
52:2305–2312. doi:
10.1128/AAC.00262-08
74. Canto
´
n E, Pema
´
n J, Sastre M et al
(2006) Killing kinetics of caspofungin,
micafungin, and amphotericin B
against Candida guilliermondii.
Antimicrob Agents Chemother
50:2829–2832. doi:
10.1128/AAC.00524-06
75. Kabbara N, Lacroix C, Peffault de
Latour R et al (2008) Breakthrough C.
parapsilosis and C. guilliermondii
blood stream infections in allogeneic
hematopoietic stem cell transplant
recipients receiving long-term
caspofungin therapy. Haematologica
93:639–640. doi:
10.3324/haematol.11149
76. Pfeiffer CD, Garcia-Effron G, Zaas
AK et al (2010) Breakthrough invasive
candidiasis in patients on micafungin.
J Clin Microbiol 48:2373–2380
77. Anderson JB (2005) Evolution of
antifungal-drug resistance:
mechanisms and pathogen fitness. Nat
Rev Microbiol 3:547–556. doi:
10.1038/nrmicro1179
78. Marco F, Lockhart SR, Pfaller MA
et al (1999) Elucidating the origins of
nosocomial infections with Candida
albicans by DNA fingerprinting with
the complex probe Ca3. J Clin
Microbiol 37:2817–2828
79. Ste
´
phan F, Bah MS, Desterke C et al
(2002) Molecular diversity and routes
of colonization of Candida albicans in
a surgical intensive care unit, as
studied using microsatellite markers.
Clin Infect Dis 35:1477–1483. doi:
10.1086/344648
80. Pfaller MA, Castanheira M, Lockhart
SR et al (2012) Frequency of
decreased susceptibility and resistance
to echinocandins among fluconazole-
resistant bloodstream isolates of
Candida glabrata. J Clin Microbiol
50:1199–1203. doi:
10.1128/JCM.06112-11
81. Cleveland AA, Farley MM, Harrison
LH et al (2012) Changes in incidence
and antifungal drug resistance in
candidemia: results from population-
based laboratory surveillance in
Atlanta and Baltimore, 2008–2011.
Clin Infect Dis 55:1352–1361. doi:
10.1093/cid/cis697
82. Garcia-Effron G, Chua DJ, Tomada JR
et al (2010) Novel FKS mutations
associated with echinocandin
resistance in Candida species.
Antimicrob Agents Chemother
54:2225–2227. doi:
10.1128/AAC.00998-09
83. Garcia-Effron G, Kontoyiannis DP,
Lewis RE, Perlin DS (2008)
Caspofungin-resistant Candida
tropicalis strains causing breakthrough
Fungemia in patients at high risk for
hematologic malignancies. Antimicrob
Agents Chemother 52:4181–4183. doi:
10.1128/AAC.00802-08
84. Hakki M, Staab JF, Marr KA (2006)
Emergence of a Candida krusei Isolate
with reduced susceptibility to
Caspofungin during therapy.
Antimicrob Agents Chemother
50:2522–2524. doi:
10.1128/AAC.00148-06
85. Fekkar A, Meyer I, Brossas JY et al
(2013) Rapid emergence of
echinocandin resistance during
Candida kefyr fungemia treatment
with caspofungin. Antimicrob Agents
Chemother 57:2380–2382. doi:
10.1128/AAC.02037-12
86. Pfaller MA (2012) Antifungal drug
resistance: mechanisms,
epidemiology, and consequences for
treatment. Am J Med 125:S3–S13.
doi:10.1016/j.amjmed.2011.11.001
87. Bizerra FC, Jimenez-Ortigosa C,
Souza ACR et al (2014) Breakthrough
candidemia due to multidrug resistant
C. glabrata during prophylaxis with
low dose of micafungin. Antimicrob
Agents Chemother 58:2438–2440. doi:
10.1128/AAC.02189-13
88. Chan TSY, Gill H, Hwang Y–Y et al
(2014) Breakthrough invasive fungal
diseases during echinocandin
treatment in high-risk hospitalized
hematologic patients. Ann Hematol
93:493–498. doi:
10.1007/s00277-013-1882-2
89. Sun H-Y, Singh N (2010)
Characterisation of breakthrough
invasive mycoses in echinocandin
recipients: an evidence-based review.
Int J Antimicrob Agents 35:211–218.
doi:10.1016/j.ijantimicag.2009.09.020
90. Myoken Y, Kyo T, Sugata T et al
(2006) Breakthrough fungemia caused
by fluconazole-resistant Candida
albicans with decreased susceptibility
to voriconazole in patients with
hematologic malignancies.
Haematologica 91:287–288
91. Beyda ND, John J, Kilic A et al (2014)
FKS mutant Candida glabrata; risk
factors and outcomes in patients with
candidemia. Clin Infect Dis. doi:
10.1093/cid/ciu407
92. Lackner M, Tscherner M, Schaller M
et al (2014) Positions and numbers of
FKS mutations in Candida albicans
selectively influence in vitro and
in vivo susceptibilities to echinocandin
treatment. Antimicrob Agents
Chemother 58:3626–3635. doi:
10.1128/AAC.00123-14
93. Liu W, Tan J, Sun J et al (2014)
Invasive candidiasis in intensive care
units in China: in vitro antifungal
susceptibility in the China-SCAN
study. J Antimicrob Chemother
69:162–167. doi:10.1093/jac/dkt330
94. Chapeland-Leclerc F, Hennequin C,
Papon N et al (2010) Acquisition of
flucytosine, azole, and caspofungin
resistance in Candida glabrata
bloodstream isolates serially obtained
from a hematopoietic stem cell
transplant recipient. Antimicrob
Agents Chemother 54:1360–1362. doi:
10.1128/AAC.01138-09
95. Pfaller MA, Diekema DJ (2007)
Epidemiology of invasive candidiasis:
a persistent public health problem.
Clin Microbiol Rev 20:133–163. doi:
10.1128/CMR.00029-06
96. Yang Y-L, Li S-Y, Cheng H–H et al
(2005) The trend of susceptibilities to
amphotericin B and fluconazole of
Candida species from 1999 to 2002 in
Taiwan. BMC Infect Dis 5:99. doi:
10.1186/1471-2334-5-99
97. Kontoyiannis DP, Lewis RE (2002)
Antifungal drug resistance of
pathogenic fungi. Lancet
359:1135–1144. doi:
10.1016/S0140-6736(02)08162-X
98. Silva S, Negri M, Henriques M et al
(2012) Candida glabrata, Candida
parapsilosis and Candida tropicalis:
biology, epidemiology, pathogenicity
and antifungal resistance. FEMS
Microbiol Rev 36:288–305. doi:
10.1111/j.1574-6976.2011.00278.x
99. Segal BH, Herbrecht R, Stevens DA
et al (2008) Defining responses to
therapy and study outcomes in clinical
trials of invasive fungal diseases:
Mycoses Study Group and European
Organization for Research and
Treatment of Cancer consensus
criteria. Clin Infect Dis 47:674–683.
doi:10.1086/590566
100. Walsh TJ, Finberg RW, Arndt C et al
(1999) Liposomal amphotericin B for
empirical therapy in patients with
persistent fever and neutropenia.
N Engl J Med 340:764–771. doi:
10.1056/NEJM199903113401004
101. Moen MD, Lyseng-Williamson KA,
Scott LJ (2009) Liposomal
amphotericin B. Drugs 69:361–392.
doi:
10.2165/00003495-200969030-00010
102. Andes DR, Safdar N, Baddley JW et al
(2012) Impact of treatment strategy on
outcomes in patients with candidemia
and other forms of invasive
candidiasis: a patient-level
quantitative review of randomized
trials. Clin Infect Dis 54:1110–1122.
doi:10.1093/cid/cis021
103. Puig-Asensio M, Pema
´
n J, Zaragoza R
et al (2014) Impact of therapeutic
strategies on the prognosis of
candidemia in the ICU. Crit Care Med.
doi:10.1097/CCM.0000000000000221
104. Cornely OA, Bassetti M, Calandra T
et al (2012) ESCMID guideline for the
diagnosis and management of Candida
diseases 2012: non-neutropenic adult
patients. Clin Microbiol Infect
18(Suppl 7):19–37. doi:
10.1111/1469-0691.12039
105. Pfaller MA, Messer SA, Boyken L
et al (2002) In vitro activities of
5-fluorocytosine against 8,803 clinical
isolates of Candida spp.: global
assessment of primary resistance using
national committee for clinical
laboratory standards susceptibility
testing methods. Antimicrob Agents
Chemother 46:3518–3521
106. Felton T, Troke PF, Hope WW (2014)
Tissue penetration of antifungal
agents. Clin Microbiol Rev 27:68–88.
doi:10.1128/CMR.00046-13
107. Glo
¨
ckner A, Cornely OA (2013)
Practical considerations on current
guidelines for the management of non-
neutropenic adult patients with
candidaemia: practical considerations
on current guidelines. Mycoses
56:11–20. doi:
10.1111/j.1439-0507.2012.02208.x
108. Shields RK, Nguyen MH, Press EG
et al (2012) The presence of an FKS
mutation rather than MIC is an
independent risk factor for failure of
echinocandin therapy among patients
with invasive candidiasis due to
Candida glabrata. Antimicrob Agents
Chemother 56:4862–4869. doi:
10.1128/AAC.00027-12
109. Baixench M-T, Aoun N, Desnos-
Ollivier M et al (2007) Acquired
resistance to echinocandins in
Candida albicans: case report and
review. J Antimicrob Chemother
59:1076–1083. doi:
10.1093/jac/dkm095
110. Bourgeois N, Laurens C, Bertout S
et al (2014) Assessment of
caspofungin susceptibility of Candida
glabrata by the Etest
"
, CLSI, and
EUCAST methods, and detection of
FKS1 and FKS2 mutations. Eur J Clin
Microbiol Infect Dis. doi:
10.1007/s10096-014-2069-z
111. Cohen Y, Karoubi P, Adrie C et al
(2010) Early prediction of Candida
glabrata fungemia in nonneutropenic
critically ill patients. Crit Care Med
38:826–830. doi:
10.1097/CCM.0b013e3181cc4734
112. Ruhnke M, Rickerts V, Cornely OA
et al (2011) Diagnosis and therapy of
Candida infections: joint
recommendations of the German
Speaking Mycological Society and the
Paul-Ehrlich-Society for
Chemotherapy. Mycoses 54:279–310.
doi:
10.1111/j.1439-0507.2011.02040.x
113. Colombo AL, Guimara
˜
es T, Camargo
LFA et al (2013) Brazilian guidelines
for the management of candidiasis—a
joint meeting report of three medical
societies: Sociedade Brasileira de
Infectologia, Sociedade Paulista de
Infectologia and Sociedade Brasileira
de Medicina Tropical. Braz J Infect
Dis 17:283–312. doi:
10.1016/j.bjid.2013.02.001
114. Bow EJ, Evans G, Fuller J et al (2010)
Canadian clinical practice guidelines
for invasive candidiasis in adults. Can
J Infect Dis Med Microbiol 21:e122–
e150
115. Castagna L, Bramanti S, Sarina B et al
(2012) ECIL 3-2009 update guidelines
for antifungal management. Bone
Marrow Transpl 47:866
116. Pappas PG, Kauffman CA, Andes D
et al (2009) Candida-clinical practice
guidelines for the management of
candidiasis: 2009 update by the
Infectious Diseases Society of
America. Clin Infect Dis 48:503–535.
doi:10.1086/596757
117. Leroux S, Ullmann AJ (2013)
Management and diagnostic
guidelines for fungal diseases in
infectious diseases and clinical
microbiology: critical appraisal. Clin
Microbiol Infect 19:1115–1121. doi:
10.1111/1469-0691.12426
118. Deshpande A, Gaur S, Bal AM (2013)
Candidaemia in the non-neutropenic
patient: a critique of the guidelines. Int
J Antimicrob Agents 42:294–300. doi:
10.1016/j.ijantimicag.2013.06.005
119. Bassetti M, Righi E, Ansaldi F et al
(2014) A multicenter study of septic
shock due to candidemia: outcomes
and predictors of mortality. Intensive
Care Med 40:839–845. doi:
10.1007/s00134-014-3310-z
120. Mermel LA, Allon M, Bouza E et al
(2009) Clinical practice guidelines for
the diagnosis and management of
intravascular catheter-related
infection: 2009 update by the
Infectious Diseases Society of
America. Clin Infect Dis 49:1–45. doi:
10.1086/599376
