Antifungal Resistance in Non‐fumigatus Aspergillus Species

ABSTRACT

This review provides an in‐depth exploration of antifungal resistance in non‐fumigatus Aspergillus species, mainly focusing on acquired resistance. The available data have been compiled and sometimes re‐analysed. It highlights the increasing prevalence of resistance in non‐fumigatus species belonging to Flavi, Terrei, Nigri, and Nidulantes Aspergillus sections, offering a detailed analysis of resistance detection methods and the global distribution of resistant strains. The review also thoroughly examines the molecular mechanisms behind resistance and raises key unresolved issues, such as the factors contributing to resistance selection and the clinical implications of in vitro resistance. Additionally, it addresses the challenges of treating infections caused by resistant Aspergillus species and cryptic species and discusses current and future strategies relying on combination therapy and newly developed antifungals. The conclusion emphasises the need for further research into resistance mechanisms and alternative treatments to address the rising threat of antifungal resistance in Aspergillus species.

1. Introduction

Acquired antifungal resistance has become an increasingly concerning issue in the field of medical mycology, particularly regarding species of Candida [1], Aspergillus [2] and dermatophytes [3]. Among Aspergillus species, while Aspergillus fumigatus is the most common species and the primary focus of research and clinical attention, in recent years, other non‐fumigatus Aspergillus species have also emerged as significant pathogens [4]. These non‐fumigatus species, which include Aspergillus flavus, Aspergillus terreus, Aspergillus niger, and Aspergillus nidulans, present unique challenges due to their inherent resistance to certain antifungal drugs [5, 6] and more limited treatment options [4]. Although acquired resistance has been mainly reported and studied in A. fumigatus , it has also emerged in these non‐fumigatus Aspergillus species [7].

As for A. fumigatus , the rise of acquired antifungal resistance in non‐fumigatus Aspergillus species is probably a multifaceted problem influenced by factors such as environmental exposure, host factors, and the inherent genetic diversity of these fungi [8]. As healthcare providers increasingly encounter infections caused by these species, understanding the mechanisms driving acquired antifungal resistance in these major human pathogens and their implications for patient care is of paramount importance.

In this review, we aimed at exploring the issues of acquired antifungal resistance in non‐fumigatus species of Aspergillus, delving into the contributing factors, resistance mechanisms, clinical implications, and potential strategies for mitigating this growing threat.

2. Materials and Methods

2.1. Literature Search Strategy

We performed database searches on PubMed, by searching studies specifically addressing resistance in non‐fumigatus Aspergillus species (e.g., A. flavus , A. terreus , A. niger , and A. nidulans ). Articles primarily focusing on A. fumigatus , non‐peer‐reviewed articles, and articles not available in full text were excluded from the review.

2.2. Data Collection and Extraction

For each study included in the review, relevant data were systematically collected and extracted. Available CYP51 nucleotide sequences for the main non‐fumigatus species were retrieved from Genbank and Ensembl Fungi databases.

2.3. Data Analysis

For molecular mechanisms of resistance, we aligned all CYP51 sequences—including introns—using the BioEdit software against the sequences of the type strains. Then, we determined the nucleotide substitutions. The sequences were then translated into proteins. For A. flavus , we considered the positions of the introns and exons published by Lucio et al. [9]. Then, we performed a comparison of protein sequences with that of strain NRRL3357. The amino acid positions shown in the figures are therefore not necessarily the same as in the published articles, since no consensus has been reached on the positions of the exons and introns in the different CYP51s. Moreover, we downloaded Cyp51A protein sequences for 86 different species from the Ensembl Fungi database. We used the mutations described in A. fumigatus to identify critical motifs that could be the sites of mutations responsible for resistance in other species.

2.4. Ethics Statement

The authors confirm that the ethical policies of the journal, as noted on the journal's author guidelines page, have been adhered to. No ethical approval was required as the research in this article is a review of the literature.

3. Results

3.1. Innate Resistance in Aspergillus spp.

Aspergillus fumigatus does not display intrinsic resistance and is generally susceptible to azoles, echinocandins, and amphotericin B [10]. However, in Aspergillus spp. both intrinsic and acquired resistance may be observed. Intrinsic resistance varies significantly across different Aspergillus species, including within a single section, but also between antifungal agents for a single species. A. flavus , which is typically susceptible to azoles and echinocandins, displays variable susceptibility to amphotericin B, with some strains displaying reduced susceptibility, though not absolute intrinsic resistance [11]. A. terreus is often considered intrinsically resistant to amphotericin B [12]. Globally, intrinsic resistance is more commonly observed in cryptic species [13], as exemplified within the Fumigati section where species such as A. lentulus [14, 15], A. udagawae [16], or A. felis [17, 18] exhibit intrinsic resistance to azoles and/or amphotericin B.

In other sections, cryptic species with intrinsic resistance include—but are not limited to—A. calidoustus in the Usti section that is resistant to azoles [19], A. alliaceus in the Flavi section that is resistant to amphotericin B and echinocandins [20], A. quadrilineatus in the Nidulantes section that is resistant to amphotericin B [21], A. tubingensis in the Nigri section that is resistant to itraconazole and/or voriconazole [22], and A. sydowii in the Versicolores section that has been reported as resistant to amphotericin B [23], although a recent study did not confirm these data [24].

In this review, we will focus on acquired resistance reported in non‐fumigatus Aspergillus species, mainly A. flavus and A. terreus , A. niger , and A. nidulans .

3.2. Methods Available for the Detection of Acquired Resistance in Non‐fumigatus Species of Aspergillus

Detecting acquired resistance is crucial for guiding effective antifungal therapy and managing fungal infections. Various laboratory techniques can be employed in clinical and environmental isolates [25]. Standardised protocols for broth microdilution assays have been developed by the Clinical and Laboratory Standards Institute (CLSI) [26] and the European Committee on Antimicrobial Susceptibility Testing (EUCAST) [27]. Clinical breakpoints and epidemiological cut‐offs/epidemiological cut‐off values (ECOFFs/ECVs) for non‐fumigatus Aspergillus species are under development thanks to ongoing efforts by both EUCAST and CLSI to provide more specific and comprehensive data. These values are crucial for detecting resistance, guiding appropriate antifungal therapy, and improving patient outcomes. ECOFFs, which are used to differentiate wild‐type strains (those without acquired resistance mechanisms) from non‐wild‐type strains (those with acquired resistance mechanisms), have been determined by EUCAST for several Aspergillus species [28]. In addition, species‐specific clinical breakpoints have been partially established for A. flavus , A. nidulans , A. niger , and A. terreus , for azole antifungals [28]. Besides reference methods, different commercially available techniques have also been developed for antifungal susceptibility testing of Aspergillus species [29]. Out of these, gradient concentration strips are the most used in clinical labs [30]. Of note, technique‐specific ECOFFs have been determined for gradient concentrations strips [31, 32], and other commercial methods, which can help to interpret MIC data in the clinical laboratory [31].

In addition to the methods described above that aimed at MIC determination, the agar screening test is a simple and effective approach that can be used for rapid assessment of azole and echinocandin susceptibility of Aspergillus species [33]. This approach is based on the incorporation of antifungal agents directly into agar plates. Once inoculated, plates are incubated and the susceptibility of Aspergillus isolates is determined based on their growth patterns [34]. This method is particularly useful for the rapid screening of azole resistance, which can then be subjected to standard MIC determination. Another advantage of this technique is that conidial suspension filtration and inoculum adjustment may not be needed [35].

Culture‐independent methods which are based on the molecular detection of resistance markers may offer a rapid and precise alternative to standard antifungal susceptibility testing of Aspergillus [36]. However, despite the availability of commercial PCR assays allowing the detection of azole resistance in A. fumigatus directly from clinical samples, this approach has not really gained interest in laboratories. Indeed, these assays yet detect only a handful of mutations associated with resistance and are not applicable to other Aspergillus species. Nevertheless, it is likely that the DNA‐based detection of antifungal resistance strategy will be increasingly used in the near future with the advance of whole‐genome sequencing along with the development of specific bioinformatic pipelines.

3.3. Reported Cases and Geographic Localisation of Acquired Resistance

3.3.1. Aspergillus Section Flavi

3.3.1.1. Azoles

Resistant strains of A. flavus have been reported in Asia, America, Europe, and Africa, including China, India, Korea, Vietnam, Taiwan, Iran, Spain, the Netherlands, Italy, France, Brazil, Argentina, Sudan, and Tunisia [9, 37, 38, 39, 40, 41, 42, 43, 44, 45, 46, 47, 48, 49]. The first report dates from 2008, when Krishnan‐Natesan et al. generated resistant strains following in vitro exposure to voriconazole [50]. The first clinical voriconazole‐resistant A. flavus strain was reported in 2012 from a lung biopsy of a Chinese patient [51]. Of 452 clinical strains isolated from the external auditory canal between April 2020 and January 2023 from the Department of Otolaryngology‐Head and Neck Surgery in central China, Peng et al. identified 92 as A. flavus , of which 17 (18.5%) strains were resistant to itraconazole (MIC > 1 mg L⁻¹), 13 (14.1%) had a MIC of 2 mg L⁻¹, 2 (2.1%) had a MIC of 4 mg L⁻¹, and one (1.1%) had a MIC of 8 mg L⁻¹ [52]. Two and five clinical azole non‐wild‐type strains were reported from India in 2015 and 2018, respectively. Four strains were non‐wild‐type for voriconazole alone, two were resistant to voriconazole and itraconazole, and one was resistant to three drugs (voriconazole, itraconazole, and posaconazole) [43, 44, 46]. More recently, in 2023, a study on fungal rhinosinusitis found that A. flavus was the main species involved in India (40/69 cases), of which 3/40 (7.5%) were resistant to itraconazole [53]. In a Korean hospital, seven resistant strains were reported in 2019 in a series of 50 clinical strains [39]. From the Mekong Delta region of Vietnam in 2020, 30/35 (85.7%) of environmental strains of A. flavus were non‐wild type for at least one antifungal among itraconazole, posaconazole, or voriconazole. Among these 30 strains, 17, 6, and 27 were non‐wild type to itraconazole, voriconazole, and posaconazole, respectively. Of particular interest, one strain has a MIC of 8 mg L⁻¹ for itraconazole [40]. The Taiwan Surveillance of Antimicrobial Resistance of Moulds (TSARM) showed that A. flavus represented 158/492 (32.1%) of collected strains and was the predominant species from clinical samples, of which 3/158 (1.9%) were resistant [47]. In Iran, Khodavaisy et al. reported on clinical and environmental strains displaying itraconazole and voriconazole MIC above ECV [54]. In another study, Zaini et al. found 4/21 (19%) non‐wild‐type isolates for voriconazole (MIC, 2–16 mg L⁻¹) with 1/21 (4.8%) resistant (MIC = 16 mg L⁻¹) [48]. In Brazil in 2017, 3/40 (7.5%) resistant strains with itraconazole MIC ≥ 4 mg L⁻¹ were reported from an environmental strains collection, while no azole‐resistant strains were found in 20 clinical strains [37]. An Argentinian study, published the same year, reported 6/18 (33.3%) environmental strains non‐wild‐type for at least one azole antifungal, while none of the 17 clinical strains showed resistance. One environmental strain displayed high MIC values to both itraconazole (≥ 8 mg L⁻¹) and voriconazole (≥ 8 mg L⁻¹) [42]. In Spain, three clinical strains resistant to azole antifungals were reported in 2020 [9]. In 2021, a Dutch team reported nine resistant strains sequentially isolated in one patient [38]. In Italy, Franconi et al. reported MICs above ECV of 0.5%, 2.1%, and 0.6% for voriconazole, posaconazole, and itraconazole, respectively, in 199 strains of A. flavus [55]. In 2023, our team reported 2/120 (1.7%) voriconazole‐ and isavuconazole‐resistant strains—from a clinical collection in two French hospitals [56]. In 2024, we identified two additional strains, resistant to both posaconazole and itraconazole [57]. A. flavus was identified as the predominant (89.9%) causative agent of fungal rhinosinusitis in Sudan. 3/88 (3.4%), 2/88 (2.3%), were non‐wild‐type to voriconazole and posaconazole, respectively. 2/88 (2.3%) and (6/88) 6.8% were resistant to itraconazole and isavuconazole, respectively [49]. Eventually, in Tunisia, out of strains from 14 haematology patients, 2/34 (5.9%) were non‐wild‐type for itraconazole and posaconazole, and 3/34 (8.8%) and 1/34 (2.9%) were non‐wild‐type only for itraconazole and posaconazole, respectively [41]. The list of A. flavus clinical resistant strains, their resistance phenotype, and the clinical context are shown in Table 1. Taken together, these data suggest that azole resistance in A. flavus is widespread worldwide and should now be monitored in clinical settings.

TABLE 1.

List of A. flavus resistant clinical strains reported in the literature.

Country (reference)	Sex/Age	Strain	Underlying diseases	Infection/Sample	Previous antifungal exposure	Treatment	Outcome	MIC (mg L−1)
								VRZ	ISA	ITZ	PSZ	AMB
China [51]	F/14	BMU29791	AML	IA/Lung	VRZ	Surgery	Cured	8	NA	2	0.25	1
India [43]	NA	NCCPF 761157	COPD	NA/Sputum	NA	NA	NA	4 (8)	NA	16	0.25	2
	50	NCCPF 760815	NA	GFR/Nasal	NA	NA	NA	2	NA	1	0.5	4
India [44]	M/33	NCCPF761476	BDB	ABPA/Sputum	VRZ and ITZ	ITZ	Discharge	8	NA	16	0.125	NA
	M/33	NCCPF761488	BDB	ABPA/Sputum	VRZ and ITZ	ITZ	Discharge	4	NA	0.5	0.125	NA
India [46]	NA	VPCI 195/P/10	COPD & NP	NA	VRZ	NA	NA	2	NA	1	0.125	NA
	NA	VPCI 982/P/15	NA	NA	NA	NA	NA	> 16	NA	> 16	> 8	NA
	NA	VPCI 985/P/15	NA	NA	NA	NA	NA	2	NA	0.25	0.06	NA
Korea [39]	F/45	E1	OM	Ear	No	NA	NA	8	NA	2	0.5	4
	F/54	E2	OM	Ear	No	NA	NA	8	NA	1	0.5	1
	F/30	E3	Cholesteatoma	Ear	No	NA	NA	8	NA	2	2	2
	M/57	E4	OM	Ear	No	NA	NA	4	NA	2	2	0.5
	M/79	E5	DM; OM	Ear	No	NA	NA	4	NA	1	0.25	1
	F/54	E6	OM	Ear	No	NA	NA	4	NA	0.5	0.25	1
	M/79	R1	COPD	Respiratory	No	NA	NA	4	NA	1	0.35	1
Spain [9, 45]	NA	CM7668	NA	NA	NA	NA	NA	0.25	0.5	> 8	1 (2)	NA
	NA	CM8087	NA	NA	NA	NA	NA	8	8	4 (8)	0.50	NA
	NA	CM9174	NA	NA	NA	NA	NA	01 (4)	2 (4)	1	0.25	NA
	NA	CM9326	NA	NA	NA	NA	NA	8	> 8	4 (8)	1 (2)	NA
	NA	CM9684	NA	NA	NA	NA	NA	> 8	2	1	0.25	NA
	NA	TP968	NA	NA	NA	NA	NA	4	4	2	1	NA
	NA	TP1004	NA	NA	NA	NA	NA	4	4	1	0.5	NA
	NA	TP1115	NA	NA	NA	NA	NA	4	4	1	0.5	NA
	NA	TP1179	NA	NA	NA	NA	NA	4	4	1	0.5	NA
Argentina [42]	NA	C2	NA	Sputum	ITZ	NA	NA	0.5	NA	0.5	0.25	≥ 8
	NA	C4	NA	Sputum	ITZ	NA	NA	0.5	NA	0.5	0.5	≥ 8
	NA	C6	NA	Sputum	VRZ	NA	NA	0.5	NA	0.5	0.5	≥ 8
	NA	C7	NA	Sputum	VRZ	NA	NA	0.5	NA	0.25	0.25	≥ 8
	NA	C11	NA	Sputum	VRZ	NA	NA	2	NA	0.25	0.25	2
	NA	C16	NA	Sputum	VRZ	NA	NA	2	NA	0.5	0.25	2
	NA	C17	NA	Sputum	VRZ	NA	NA	2	NA	0.5	0.5	1
The Netherlands [38]	F/66	V‐152‐49	COPD	NA	NA	NA	NA	4	NA	1	0.25	2
	F/66	V‐156‐58	COPD	NA	VRZ	VRZ	Dead	> 16	NA	16	0.5	2
	F/66	V‐158‐11	COPD	NA	VRZ	VRZ	Dead	> 16	NA	> 16	1	2
	F/66	V‐158‐20	COPD	NA	VRZ	VRZ	Dead	16	NA	16	0.5	1
	F/66	V‐158‐70	COPD	NA	VRZ	VRZ	Dead	> 16	NA	2	0.5	2
	F/66	V‐158‐75	COPD	NA	VRZ	VRZ	Dead	> 16	NA	> 16	1	2
	F/66	V‐158‐76	COPD	NA	VRZ	VRZ	Dead	> 16	NA	> 16	1	2
	F/66	V‐158‐77	COPD	NA	VRZ	VRZ	Dead	16	NA	1	0.25	2
	F/66	V‐159‐56	COPD	NA	VRZ	VRZ	Dead	> 16	NA	> 16	2	2
	F/66	V‐159‐40	COPD	NA	VRZ	VRZ	Dead	> 16	NA	1	0.5	2
France [56, 57]	M/27	HEGP 2885	Lung Tx	Colonisation/NA	VRZ	NA	NA	8	8	1	1	4
	45	HEGP 3517	Kidney cancer	IA/NA	VRZ	NA	NA	8	8	2	1	2
	NA	GRE 150	NA	NA	NA	NA	NA	0.5	0.5	> 126	64	NA
	22	LIL 226	CF	ABPA//NA	ITZ	NA	NA	0.5	1	> 126	> 126	NA
	M/48	HEGP 362	NA	NA	NA	NA	NA	0.5	1	0.25	0.5	≥ 8
Taiwan [47]	NA	2020‐C01‐010	NA	Wound	NA	NA	NA	4	1	0.5	0.25	NA
	NA	2020‐C03‐025	NA	Sputum	NA	NA	NA	> 16	16	1	0.5	NA
	NA	2020‐S07‐003	NA	Ear	NA	NA	NA	8	4	1	0.25	NA
Sudan [49]	NA	v313‐29	NA	Rhinosinusitis	NA	NA	NA	2	8	2	1	NA
	NA	M.064–11	NA	Rhinosinusitis	NA	NA	NA	4	2	0.25	0.125	NA
	NA	M.064–12	NA	Rhinosinusitis	NA	NA	NA	4	2	0.25	0.25	NA
	NA	M.064–28	NA	Rhinosinusitis	NA	NA	NA	4	4	16	8	NA
Tunisia a [41]	NA	TN‐7	HC	IA/nasal	NA	NA	NA	NA	NA	1.5	0.125	NA
	NA	TN‐15	HC	IA/nasal	NA	NA	NA	NA	NA	1	0.19	NA
	NA	TN‐16	HC	IA/sputum	NA	NA	NA	NA	NA	1	0.19	NA
	NA	TN‐31	HC	IA/lung biopsy	NA	NA	NA	NA	NA	1.5	0.75	NA
	NA	TN‐32	HC	IA/nasal	NA	NA	NA	NA	NA	0.5	1	NA
	NA	TN‐33	HC	IA/nasal	NA	NA	NA	NA	NA	1	0.75	NA

Abbreviations: ABPA, allergic broncho pulmonary aspergillosis; AMB, amphotericin B; AML, acute myeloid leukaemia; BDB, basic disease bullous lung; CF, cystic fibrosis; COPD, chronic obstructive pulmonary disease; DM, Diabetes Mellitus; GFR, granulomatous fungal rhinosinusitis; HC, haematological cancer; IA, invasive aspergillosis; ISA, isavuconazole; ITZ, itraconazole; MIC, minimal inhibitory concentration; NA, not available; NP, nasal polyposis; OM, otitis media; PSZ, posaconazole; Tx, transplant; VRZ, voriconazole.

^a Antifungal susceptibility determined with the Etest; other MICs were determined by CLSI or EUCAST method. For MICs, the second values in brackets correspond to the value of the replicate, when the second value is different from the first.

3.3.1.2. Amphotericin B

Species in section Flavi are known to have high MICs to amphotericin B but are not considered to have intrinsic resistance, except for A. alliaceus, which always has an amphotericin B MIC of ≥ 8 mg L⁻¹ [20, 58, 59, 60].

Based on a literature review that included data from 3663 strains, Fakhim et al. estimated resistance to amphotericin B in A. flavus at 14.9%. In Europe and the USA, these resistance rates were estimated to be 14.3% and 11.7%, respectively [61]. Interpretation of these data remains difficult as different breakpoints may have been used over the years and across the different publications. Indeed, clinical breakpoints for amphotericin B against A. flavus are not yet available, but EUCAST has set an ECOFF at ≤ 4 mg L⁻¹. In a previous publication, we observed that 83/120 (69.2%) of the strains from a French hospital had a MIC ≥ 2 mg L⁻¹ [56]. In the same study, 1/120 (0.8%) with a MIC of 8 mg L⁻¹ was considered to have acquired resistance. In Argentina, Hermida et al. reported 4/17 (23.5%) clinical strains with MICs ≥ 8 mg L⁻¹ [42]. In Spain, Lucio et al. also reported a clinical strain resistant to amphotericin B (MIC in double determination at 8 and > 16 mg L⁻¹) [9]. Eventually, Franconi et al. reported 3% of amphotericin B MICs above ECV in a collection of 199 strains of A. flavus from Italy [55].

3.3.1.3. Echinocandins

Compared with azoles, there are few data for echinocandins. In Iran, 6/14 (42.9%) A. flavus clinical strains with MICs of 4 mg L⁻¹ or above have been reported [62]. One A. flavus isolate with caspofungin MIC > 32 mg L⁻¹, isolated simultaneously with A. fumigatus , caused a fatal infection in a heart‐transplant patient [63].

3.3.2. Aspergillus Section Terrei

3.3.2.1. Azoles

Non‐wild‐type but also azole‐resistant strains of A. terreus have been reported from both clinical and environmental samples. The first description of clinical azole‐resistant isolates dates from the early 2000s in Spain when Gomez‐Lopez and colleagues evidenced strains with itraconazole and voriconazole MICs up to 8 and 4 mg L⁻¹, respectively [64]. Subsequently, two studies focusing on CF patients in Denmark reported one patient with an itraconazole‐ and voriconazole‐resistant isolate [65] and successive pan‐azole‐resistant isolates from the same patient from 2003 to 2011 after long‐term exposure to azoles [66]. In France, 1/61 (1.6%) A. terreus clinical isolates exhibited pan‐azole resistance, while 3/61 (4.9%) were resistant to at least one azole drug [67]. In Austria, two studies reported azole‐resistant isolates in both clinical and environmental samples [68, 69]. In the first study that included isolates collected between 1996 and 2006, the maximum MIC was 4 mg L⁻¹ against voriconazole for both clinical and environmental isolates, which is above ECOFF (2 mg L⁻¹), and the maximum MIC against posaconazole was 0.5 mg L⁻¹ and 0.25 mg L⁻¹ for clinical and environmental isolates, respectively (clinical breakpoint: 0.25 mg L⁻¹) [69]. In their last study, which included isolates collected during 2019–2020, voriconazole and posaconazole resistance was observed in 3/51 (5.9%) and 5/51 (9.9%) of clinical isolates, respectively, but posaconazole resistance reached 54/238 (22.6%) in environmental isolates [68]. In Portugal, over a 4‐year period, 1/27 (3.7%) environmental isolates was itraconazole‐resistant, but no resistance was detected in clinical isolates (n = 13) [70]. Screening of a large worldwide collection of strains (n = 432) isolated during 2014–15 from 21 countries by TerrNet Study group also identified posaconazole resistance, with a rate range from 0% to 15% according to the country [71]. Posaconazole‐resistant isolates were found in Germany, UK, Austria, France, Italy, and Spain. It has to be noted that all posaconazole‐resistant isolates had moderately elevated posaconazole MICs of 0.5–1 mg L⁻¹, for a breakpoint at > 0.25 mg L⁻¹. Moreover, voriconazole resistance was rare and itraconazole resistance was absent. Another international study, investigating respiratory isolates of Terrei section, reported 1/40 (2.5%) and 8/40 (20%) isolates resistant to itraconazole and posaconazole, respectively [72]. In the US, voriconazole (1.14% of 175 isolates) and isavuconazole (6.77% of 192 isolates) non‐wild‐type isolates have been described [73]. In Martinique Island (West Indies), 2/2 (100%) clinical isolates were resistant to posaconazole and isavuconazole, of which one was non‐wild‐type for voriconazole [74]. In China, among clinical isolates collected from 1999 to 2019, 1/48 (2.1%) was resistant to itraconazole [75]. Only 1/49 (2%) isolate collected in Taiwan between 2016 and 2020 and tested against azoles was non‐wild‐type to voriconazole [47].

3.3.2.2. Echinocandins

Compared to azoles, isolates with high MECs against echinocandins are rarely reported in the literature with, to the best of our knowledge, only two isolates have been reported to date. One isolate had caspofungin MEC ≥ 8 mg L⁻¹ [76], and one isolate had caspofungin MEC = 2 mg L⁻¹ [77].

3.3.2.3. Amphotericin B

Aspergillus terreus is generally intrinsically resistant to amphotericin B, with most strains showing high MIC of ≥ 2 mg L⁻¹. This in vitro resistance is associated with a poor outcome in patients treated with amphotericin B [78]. However, a small percentage (8%–13%) of A. terreus isolates globally exhibit lower MICs and are susceptible to amphotericin B [79].

3.3.3. Aspergillus Section Nigri

3.3.3.1. Azoles

Aspergillus niger is reported to be less common in human infections than A. tubingensis and A. welwitschiae [80, 81, 82, 83]. Aspergillus niger and A. tubingensis are sometimes present in equal proportions in epidemiological studies of the Nigri section [84, 85]. In this chapter, we will focus on these two species and other species of the Nigri section.

In Japan, both clinical and environmental strains of section Nigri ( A. niger ss, A. tubingensis, and A. welwitschiae), with MICs > 2 mg L⁻¹ for voriconazole or itraconazole, have been reported [86]. Among clinical isolates, itraconazole MICs > 2 mg L⁻¹ were observed in 1/4 (25%) A. niger , 2/4 (50%) A. welwitschiae, and 16/20 (80%) A. tubingensis. Voriconazole MICs > 2 mg L⁻¹ were found in 5/14 (35.7%) A. niger , 8/46 (17.4%) A. welwitschiae, and 18/20 (90%) A. tubingensis. Among environmental isolates, itraconazole MICs > 2 mg L⁻¹ were detected in 5/6 (83.3%) A. niger , 2/13 (15.4%) A. welwitschiae, and 15/19 (78.9%) A. tubingensis. Similarly, voriconazole MICs > 2 mg L⁻¹ were observed in 6/6 (100%) A. niger , 3/13 (23.1%) A. welwitschiae, and 17/19 (89.5%) A. tubingensis. In India, Sen et al. observed resistance in section Nigri isolates from soil samples, with 19/161 (11.8%) resistant to itraconazole, 5/161 (3.1%) resistant to posaconazole, and 19/161 (1.24%) resistant to voriconazole. Of the resistant strains, 20 were identified at the species level, revealing 13 A. niger and 7 A. tubingensis [87]. Also in India, a study on 69 cases of fungal rhinosinusitis found six cases due to A. niger , all being itraconazole‐resistant [53]. In Iran, a study was conducted on 134 clinical and environmental strains of the Nigri section, including 72 A. niger strains, 49 A. tubingensis strains, 1 A. uvarum (clinical), 1 A. acidus (soil), and 1 A. sydowii (clinical). Of 39 clinical A. niger strains, seven had itraconazole MICs > 16 mg L⁻¹, and six had voriconazole MICs > 16 mg L⁻¹. Of 33 environmental A. niger strains, six had itraconazole MICs > 16 mg L⁻¹ and six had voriconazole MICs > 16 mg L⁻¹. Of 29 environmental A. tubingensis strains, 11 had itraconazole MICs > 16 mg L⁻¹. No clinical A. tubingensis had MICs > 16 mg L⁻¹ for either itraconazole or voriconazole [88]. Of 452 clinical strains isolated from the external auditory canal between 2020 and 2023 in China, Peng et al. identified 2/55 (3.6%) A. niger strains resistant to itraconazole, both with a MIC of > 8 mg L⁻¹, and 1 strain resistant to voriconazole with a MIC of 8 mg L⁻¹ [52]. In a study evaluating the occupational exposure of Danish workers, Kofoed et al. reported azole resistance in 19/131 (14.5%) strains of the Nigri section [89]. In Italy, Latta et al. reported two azole‐resistant strains of the Nigri section colonising the respiratory tract of haematology patients. One, identified as A. tubingensis strain, had an itraconazole MIC of 64 mg L⁻¹ and a posaconazole MIC of 1 mg L⁻¹, while the other, identified as A. niger , displayed a posaconazole MIC of 1 mg L⁻¹ [90]. In a more recent study, Franconi et al. reported 0.8% of MICs above ECV for voriconazole on a large collection of A. niger strains (n = 251) [55]. In Spain, a study identified 7/36 (19.4%) clinical and environmental strains of section Nigri resistant (MIC > 4 mg L⁻¹ for itraconazole, MIC > 2 mg L⁻¹ for voriconazole) to at least one azole, including one clinical and one environmental strain of A. niger , four environmental strains of A. tubingensis, and one strain of A. brasiliensis [91]. In 2011, another study investigating 45 clinical strains of section Nigri from the Mycology Reference Centre, Manchester, UK, identified 25 (55.6%), 8 (17.8%), 6 (13.3%), 3 (6.7%), and 3 (6.7%) of A. awamori, A. tubingensis, A. niger , A. acidus, and unspecified cryptic species strains, respectively. High rates of itraconazole resistance were unveiled. Interestingly, variable rates of resistance were noted between species. Itraconazole resistance ranged from 33% in A. niger to 100% from the A. acidus group [22]. Finally, five strains of A. tubingensis responsible for invasive aspergillosis with MICs of itraconazole between 4 and 8 mg L⁻¹ were reported in Belgium [83]. Overall, these findings confirm previous statements by Hendrickx et al. and Howard et al. who observed that A. niger was more susceptible to itraconazole and voriconazole than A. tubingensis [22, 84].

3.3.3.2. Amphotericin B

In a large study conducted at Pisa University Hospital in Italy aiming at investigating the in vitro susceptibility of A. niger to amphotericin B, Franconi et al. reported 21.8% MICs above amphotericin B ECV (n = 251 strains) [55]. Overall, in the EUCAST database (https://mic.eucast.org/search/), only 0.5% of A. niger isolates had amphotericin B MIC of ≥ 4 mg L⁻¹.

3.3.4. Aspergillus Section Nidulantes

3.3.4.1. Azoles

Azole resistance is still rarely reported in A. nidulans but possibly underestimated as this species is only rarely associated with aspergillosis (MIC are therefore usually not determined). Although several epidemiological studies did not detect azole resistance [47, 72, 92, 93], there are a few outlining that resistance may also occur within this section. In one study aiming to determine wild‐type MIC distributions and ECVs for triazoles in a large collection of Aspergillus species, 2/141 (1.4%) A. nidulans isolates had itraconazole MIC above 2 mg L⁻¹ using the CLSI method [94]. More recently, another study that tested 60 clinical and environmental isolates of Aspergillus section Nidulantes from five countries found 26.8% clinical strains with itraconazole MIC higher than the current EUCAST breakpoint (MIC > 2 mg L⁻¹). Most (90.9%) belonged to A. spinulosporus whereas A. nidulans sensu stricto accounted for 9.1% [95]. Additionally, two clinical isolates of A. spinulosporus had intermediate MIC for itraconazole. Of the 13 environmental isolates tested, two were itraconazole‐resistant (one A. quadrilineatus and one A. rugulosus ). In the EUCAST database, 4%–6% of A. nidulans isolates exhibit MIC of ≥ 8 mg L⁻¹ for azoles.

3.3.4.2. Amphotericin B

It has been suggested that A. nidulans could be intrinsically resistant to amphotericin B [96] because this species has generally higher MICs to amphotericin B than A. fumigatus . In a recent study, the MIC₉₀ for amphotericin B was 32 mg L⁻¹ for A. nidulans sensu stricto and for A. spinulosporus [95].

3.3.4.3. Echinocandins

Although some echinocandins (e.g., anidulafungin) are derived from natural products synthesised by A. nidulans , these drugs are generally active against species belonging to this complex. There are, however, some variations of echinocandin susceptibility between species, with A. spinulosporus showing higher caspofungin MIC (MIC₉₀ = 8 mg L⁻¹) than A. nidulans (MIC90 = 1 mg L⁻¹) and A. quadrilineatus (MIC₉₀ = 4 mg L⁻¹). In the same study, some isolates of A. quadrilineatus had high MICs for micafungin and anidulafungin, indicating that acquired echinocandin resistance may be possible.

3.4. Molecular Mechanisms of Resistance

In this section, we will focus mainly on the mechanisms of resistance to azole antifungals, with particular emphasis on those related to CYP51, encoding the cellular target of azole drugs. Of note, unlike section Flavi, all Aspergillus sections including section Fumigati have two CYP51s genes (CYP51A and CYP51B). Both catalyse critical steps in parallel pathways [97, 98, 99]. CYP51A is involved in the main pathway which may explain why amino acid substitutions associated with resistance occur mostly in Cyp51A and have a stronger impact on azole susceptibility than those reported in Cyp51B. In the section Flavi, the homology between Cyp51C, the Cyp51A paralog, and Cyp51A suggests that Cyp51C catalyses the same pathway as Cyp51A and therefore, mutations in this gene could also contribute to azole resistance. However, basal and induced expression of CYP51B and CYP51C was less important than CYP51A expression [9, 44]. This lower expression of CYP51B and CYP51C is an additional argument that allows us to hypothesise that the main Cyp51 protein potentially involved in resistance mechanisms is CYP51A. Overall, Cyp51A protein sequences show at least 66.4% similarity in a comparison of sequences from 86 Aspergillus species from different sections. They share several motifs that are conserved across species. In A. fumigatus , Cyp51A substitutions previously reported to be associated with resistance are listed in Table 2. These amino acid changes occur at residues highly conserved between Aspergillus species. Of note, some species exhibit natural variation at these residues such as M220 motif showed also as I218 (section Nidulantes species), as L218 (A. alliaceus and A. coremiiformis), or V218 (A. calidoustus) which is likely to explain natural variations in azole susceptibility between species. Note that A. calidoustus shows intrinsic resistance to azole antifungals [59, 100].

TABLE 2.

Cyp51A amino acid positions from different Aspergillus species compared with positions known to be responsible for azole resistance in A. fumigatus when mutated.

Section	Species	GenBank accession number	Amino‐acid
Fumigati	A. fumigatus	DS499598	G54	L98	Y121	G138	P216	F219	M220	A284	T289	Y431	G432	G434	G448
Fumigati	A. fischeri	EAW25441	G54	L98	Y121	G138	P216	F219	M220	A284	T289	Y431	G432	G434	G448
Fumigati	A. lentulus	GFF61070	G54	L98	Y121	G138	P216	F219	M220	A284	T289	Y431	G432	G434	G448
Fumigati	A. novofumigatus	PKX93118	G54	L98	Y121	G138	P216	F219	M220	A284	T289	Y431	G432	G434	G448
Fumigati	A. thermomutatus	RHZ54084	G54	L110	Y133	G150	P228	F231	M232	A296	T301	Y443	G444	G446	G460
Aspergillus	A. glaucus	OJJ84218	G54	L98	Y121	G138	P216	F219	M220	A286	T291	Y434	G435	G437	G451
Aspergillus	A. cristatus	ODM18444	G54	L98	Y121	G138	P216	F219	M220	A286	T291	Y434	G435	G437	G451
Aspergillus	A. ruber	EYE96192	G54	L98	Y121	G138	P216	F219	M220	A283	T288	Y430	G431	G433	G447
Candidi	A. campestris	PKY03207	G51	L95	Y118	G135	P213	F216	M217	A288	T293	Y440	G441	G443	G457
Candidi	A. candidus	PLB41256	G51	L95	Y118	G135	P213	F216	M217	A288	T293	Y440	G441	G443	G457
Candidi	A. taichungensis	PLN81727	G52	L96	Y119	G136	P214	F217	M218	A288	T293	Y439	G440	G442	G456
Circumdati	A. steynii	PLB55559	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y425	G426	G428	G442
Clavati	A. clavatus	EAW10153	G54	L98	Y121	G138	P216	F219	M220	A284	T289	Y430	G431	G433	G447
Flavi	A. flavus	XM_002375082	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. oryzae	XP_001819419	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. alliaceus	KAB8235133	G52	L96	Y119	G136	P214	F217	L218	A280	T285	Y424	G425	G427	G441
Flavi	A. arachidicola	PIG89481	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. avenaceus	KAE8151200	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. bertholletiae	KAE8374617	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. bombycis	OGM50467	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. caelatus	KAE8360647	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. coremiiformis	KAE8354520	G52	L96	Y119	G136	P214	F217	L218	A280	T285	Y424	G425	G427	G441
Flavi	A. leporis	KAB8071199	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. minisclerotigenes	KAB8274449	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. novoparasiticus	KAB8224139	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. parasiticus	KAB8206802	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. pseudocaelatus	KAE8419072	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. pseudonomiae	KAE8409683	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. pseudotamarii	KAE8137678	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. sergii	KAE8328643	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Flavi	A. tamarii	KAE8160134	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Nidulantes	A. mulundensis	RDW81169	G52	L96	Y119	G136	P214	F217	I218	A280	T285	Y428	G429	G431	G445
Nidulantes	A. nidulans	CBF85786	G52	L96	Y119	G136	P214	F217	I218	A280	T285	Y428	G429	G431	G445
Nidulantes	A. sydowii	OJJ64317	G52	L96	Y119	G136	P214	F217	I218	A280	T285	Y429	G430	G432	G446
Nigri	A. niger	AEK81582	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Nigri	A. tubingensis	AEK81587	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Nigri	A. aculeatinus	RAH74471	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y426	G427	G429	G443
Nigri	A. aculeatus	OJK02412	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y426	G427	G429	G443
Nigri	A. brunneoviolaceus	RAH42286	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y426	G427	G429	G443
Nigri	A. awamori	GCB20302	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Nigri	A. brasiliensis	OJJ77622	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Nigri	A. ellipticus	PYH98808	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y425	G426	G428	G442
Nigri	A. eucalypticola	PWY64102	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Nigri	A. fijiensis	RAK73127	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y426	G427	G429	G443
Nigri	A. heteromorphus	PWY78270	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y425	G426	G428	G442
Nigri	A. homomorphus	RAL16372	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y426	G427	G429	G443
Nigri	A. ibericus	RAK97770	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y425	G426	G428	G442
Nigri	A. indologenus	PYI33145	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y426	G427	G429	G443
Nigri	A. japonicus	RAH79284	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y426	G427	G429	G443
Nigri	A. carbonarius	OOF99113	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y425	G426	G428	G442
Nigri	A. costaricensis	RAK85439	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Nigri	A. luchuensis	OJZ80911	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Nigri	A. neoniger	PYH34323	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Nigri	A. piperis	RAH57528	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y424	G425	G427	G441
Nigri	A. saccharolyticus	PYH45087	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y426	G427	G429	G443
Nigri	A. sclerotiicarbonarius	PYI11361	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y425	G426	G428	G442
Nigri	A. scleroniger	PWY91912	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y425	G426	G428	G442
Ochraceorosei	A. ochraceoroseus	PTU24382	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y426	G427	G429	G443
Ochraceorosei	A. rambellii	KKK26746	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y426	G427	G429	G443
Polypaecilum	A. sclerotialis	RJE21812	G52	L96	Y119	G136	P214	F217	M218	A280	T285	Y422	G423	G425	G439
Terrei	A. terreus	EAU33678	G51	L95	Y118	G135	P21 3	F216	M217	A285	T290	Y431	G4 32	G434	G448
Usti	A. calidoustus	CEL05254	G52	L96	Y119	G136	P214	F217	V218	A280	T285	Y428	G429	G431	G445

Note: Bold: acquired mutations in resistant strains already reported for this position; Red: species polymorphism; Red and bold: species polymorphism associated with intrinsic resistance.

3.4.1. Aspergillus Section Flavi

3.4.1.1. Azole Antifungals

Compilation from articles and directly from GenBank allowed us to find 153 strains of A. flavus for which at least one of the CYP51 genes was sequenced. Amino acid substitutions, resistance profile, MIC, and nucleotide GenBank accession number are presented in Table S1 and summarised in Table 3 and Figure 1.

TABLE 3.

Compilation from articles and directly from GenBank of strains for which at least one of the CYP51s was sequenced.

Reference	Susceptibility profile	Number	Amino‐acid substitution			MIC
			Cyp51A	Cyp51B	Cyp51C	VRZ	ITZ	PSZ	ISA
[38]	R	8	Y119F	NA	NA	16–> 16	1–> 16	0.25–2	NA
[39]	S	42	NA	NA	T34A‐M54T‐S240A‐R250ST‐D254N‐D254G ‐P257S‐Q275K‐I285V‐S399I‐N423D	NA	NA	NA	NA
[39]	R	7	None	None	T34A‐M54T‐S240A‐D254G‐N423D	4–8	0.5–2	0.25–0.5	NA
[56, 57]	R	6	N31Y‐ P214L‐K322N‐E410V	K165E	T34A‐M54T‐S240A‐R250ST‐D254G‐D254N‐I285V‐S399I‐N423D b	0.5–8	1–> 8	1–> 8	0.5–8
[56, 57]	S	3	None	None	M54T‐S240A‐D254G‐N423D	1	0.25–0.5	0.25–0.5	1–2
[42]	S	7	N31Y‐K322N	None	M54T ‐K230T ‐S240A ‐L266F ‐S361W‐ L366F ‐A397P‐N423D	2	0.25–1	0.125–0.5	NA
[42]	R	6	N31Y‐K322N	None	M54T‐S240A‐ G333A ‐S361W‐T385P ‐A397P‐ F403S ‐N423D‐ D426E	2–≥ 8	0.5–≥ 8	0.25–0.5	NA
[51]	R	1	None	None	M54T‐S240A‐P419T ‐N423D	8	2	NA	NA
[9]	S	12	A199T‐K322N‐T329A	K165E	M54T‐S240A‐R250ST‐D254N‐D254G‐P276T‐I285V‐S399I‐N423D	0.5–2	0.25–1	0.125–0.25	0.25–2
[9]	R	9	P214L‐ T329A	None	M54T‐S240A‐D254G‐H349R‐N423D	0.25–> 8	1–> 8	0.5–2	0.5–> 8
[43]	R	1	A199T	None	M54T‐S240A‐D254N‐I285V‐Y319H	8	16	0.25	NA
[43]	S	5	A199T	None	M54T‐S240A‐D254N‐I285V	0.25–2	0.06–1	0.03–0.5	NA
[44]	S	1	A199T	None	M54T‐S240A	0.5	0.12	0.06	NA
[44]	R	2	A199T	None	M54T‐S240A	4–8	0.5–16	0.13	NA
[46]	S	2	A199T‐R444S	Q354K	S196F‐ S240A‐D254N‐I285V ‐A324P‐ N423D‐ V465M	2	0.25–1	0.06–0.125	NA
[46]	R	1	A199T‐R444S		M54T‐S240A	> 16	> 16	> 8	NA
[47]	R	3	N31Y‐A199T‐K322N‐G441S	None	M54T‐S240A‐D254G‐N423D	4–> 16	0.5–1	0.25–0.5	1–16
[47]	S	5	N31Y‐A199T‐K322N	None	M54T‐S240A‐D254N‐D254G‐ Q275N‐ I285V‐Y319H‐N423D	NA	NA	NA	NA
Bavadharani, S a	NA	8	N31Y‐K32M‐R194L‐L197I‐T329A	L341F	T34A‐M54T‐D172N‐S240A‐K248N‐R250ST‐D254N‐D254G‐I285V‐N423D	NA	NA	NA	NA
[101]	NA	1	None	NA	NA	NA	NA	NA	NA
[41]	S	5	E200K	None	NA	NA	0.125–0.5	0.094–0.125	NA
[41]	R	6	None	K267E	NA	NA	0.5–1.5	0.125–1	NA
[49]	R	4	A199T	NA	NA	2–4	0.25–16	0.125–8	2–8
Nargesi, S a	NA	5	N31Y‐K322N	NA	NA	NA	NA	NA	NA
Brito Devoto, T a	NA	3	None	NA	NA	NA	NA	NA	NA

Note: In red: reported only in resistant strains; in blue: reported in both resistant and susceptible strains; in green: reported only in susceptible strains; black: reported in unknown phenotype.

Abbreviations: ISA, isavuconazole; ITZ, itraconazole; MIC, minimal inhibitory concentration; NA, not available; PSZ, posaconazole; R, resistant; S, susceptible; VRZ, voriconazole.

^a Unpublished only available on GenBank.

^b One strain with a deletion of CYP51C.

FIGURE 1.

Schematic representation of the A. flavus CYP51 genes showing all mutations leading to amino acid substitutions previously reported in the literature. Sequences were retrieved from Genbank. Green: substitutions only found in susceptible strains; red: reported only in resistant strains, blue: reported in both resistant and susceptible strains, black: reported in unknown phenotype. A: CYP51A, B: CYP51B, and C: CYP51C.

The CYP51A promoter showed point substitutions but no tandem repeats. In the Cyp51A, a total of 13 amino acid substitutions were found, four of which (Y119F, P214L, E410V, and G441S) are exclusively found in resistant strains [9, 38, 47, 57]. Y119F and G441S are responsible for resistance to voriconazole, and P214L is responsible for resistance to itraconazole; they correspond respectively to Y121F, G448S, and P216L in A. fumigatus [102]. Moreover, in the A. flavus laboratory‐resistant selected strain, Krishnan‐Natesan et al. described the substitutions Y132N, K197N, D282E, M288L, and T469S in Cyp51A, although their involvement in antifungal resistance remains to be determined [50]. Of note, Krishnan‐Natesan et al. consider different introns and exons than those used in recent publications. As a result, the substitution numbers in their original publication do not match those presented in the present article. As defined in recent publications, these substitutions correspond to Y66N, K131N, D276E, M282L, and T463S.

For Cyp51B, the K267E substitution has been reported in both susceptible and resistant strains, and the Q354K has been reported in a strain with an MIC of 2 mg L⁻¹ for VRZ. A third mutation (L341F) has been reported in Genbank but has not yet been published in an article, and no information on susceptibility is available. In addition, Krishnan‐Natesan et al. reported 4 mutations (H399P, D411N, T454P, and T486P) in a resistant strain selected in vitro [50]. Nevertheless, the positions of exons were different in this publication, which means that the three substitutions are more likely H422P, D434N, and T477P. The last one, T486P, is not found in the coding region considered in this article.

Cyp51C shows a more pronounced polymorphism with 38 amino acid substitutions and a replacement of arginine by a stop codon in position 250. The main amino acid substitutions only present in resistant strains are H349R and P419T [9, 51] although their involvement in azole resistance needs to be confirmed experimentally. Eventually, the Y319H substitution reported by Paul et al. [43] to be involved in resistance was found by Wang et al. in a susceptible strain [47].

Besides amino acid substitutions, CYP51 genes may be involved in azole resistance through other mechanisms, which include copy number variation of these genes or upregulation (the former scenario also conferring overexpression). In vitro, Liu et al. showed that an extra copy of CYP51A or CYP51B confers resistance to itraconazole and voriconazole, while an extra copy of CYP51C leads to itraconazole resistance only. He also reported slight overexpression of CYP51C in one voriconazole‐resistant strain [51]. Paul RA et al. showed that CYP51C expression is only detectable in non‐wild‐type isolates, except one wild‐type isolate of his panel [44]. Sharma et al. also reported overexpression of CYP51A, CYP51B, and CYP51C in one resistant strain compared to susceptible strains [46]. Lucio et al. showed an increase in expression of CYP51C in all resistant strains from his collection compared to reference strain ATCC 2024304. In the same panel, one resistant strain showed significant overexpression of CYP51A [9].

Furthermore, a complete deletion of CYP51C has been reported in a resistant strain that also has a P214L substitution in Cyp51A [57] showing that CYP51C is not essential in this species.

3.4.1.1.1. Active Efflux

As some azole‐resistant isolates failed to show mutations or overexpression of CYP51 genes, other putative mechanisms have been investigated. Efflux pumps, by preventing accumulation of antifungal drugs, can participate in antifungal resistance and have been reported in other fungal species. In this respect, exposure to subinhibitory concentrations of voriconazole led to overexpression of several ABC (ATP‐binding cassette) (MDR1‐4, ATRF) and MFS (Major Facilitator Superfamily) transporters in both voriconazole‐resistant clinical isolates and laboratory‐selected isolates [103, 104]. Basal overexpression of MDR1, MDR2, and MFS1 was observed in a clinical isolate carrying mutations in CYP51A and CYP51C genes [46]. Basal overexpression of ATRF and MFS1 but reduced expression of MDR2 was observed in seven clinical voriconazole‐resistant strains [39]. After in vitro exposure to voriconazole, the expression of CDR1B increased in two clinical isolates. One also demonstrated overexpression of MDR1, MDR2, MDR4, and ATRF [44]. Of note, the azole‐resistant strain obtained after serial in vitro cultures with voriconazole harboured a T1673G mutation (L558W) in the gene encoding the bZIP transcription factor YAP1. This mutation was associated with a 62‐fold overexpression of the ATRF transporter. Reversion of this mutation and the null mutant of ATRF led to a susceptible phenotype, suggesting that L558W may act as a gain‐of‐function [105]. Collectively, these data suggest the involvement of efflux pumps in azole resistance in A. flavus .

3.4.1.1.2. Other Mechanisms

Isolates exposed to voriconazole presented a duplication of chromosome 8, and in some cases, this was associated with a segmental duplication of chromosome 3, [106] suggesting that aneuploidy could also take part in azole resistance. Overexpression of the genes encoding aldehyde reductase (AKR1) and ABC fatty acid transporters was reported in two non‐WT isolates showing no mutations in the CYP51 genes [48].

3.4.1.2. Amphotericin B

Various mechanisms contributing to amphotericin B resistance in pathogenic fungi have been described and include decreased membrane ergosterol content, reduction of oxidative stress, and alterations of the fungal cell wall [107]. However, the mechanism explaining the reduced susceptibility of A. flavus to amphotericin B has been poorly studied. Based on what is known in A. terreus , one study explored potential mechanisms of amphotericin B resistance in A. flavus [108]. Among a collection of 117 clinical isolates, one isolate with an amphotericin B MIC of 32 mg L⁻¹ was identified. This isolate exhibited elevated levels of endogenous reactive oxygen species and hypersensitivity to oxidative stress, which contrasts with observations in A. terreus . Unfortunately, the study did not assess ergosterol membrane levels. In vitro selected A. flavus amphotericin B‐resistant isolate showed unchanged levels of ergosterol but modifications in the cell wall composition (increased in 1,3‐α glucans) that could participate in amphotericin B resistance by preventing the drug from reaching the cell membrane [109]. Overall, the results suggest that the mechanisms responsible for amphotericin B resistance in A. flavus may differ from those reported in A. terreus .

3.4.1.3. Echinocandin

Among non‐fumigatus Aspergillus, there is a paucity of data with, to the best of our knowledge, only two isolates of A. flavus shown to be echinocandin‐resistant. Of note, resistance was not linked to fks mutations [62].

3.4.2. Aspergillus Section Terrei

3.4.2.1. Azole Antifungals

As for A. flavus , there is still a paucity of data regarding the molecular mechanisms beyond acquired azole resistance in Aspergillus section Terrei [71]. Exploration of the mechanism of azole resistance of isolates from theTerrNet Study group showed that although most had a wild‐type phenotype, sequencing of the CYP51A of posaconazole‐resistant isolates identified mutations at codon 217 (M217T and M217V) in two isolates along with E319G and A221V [71]. None of these substitutions were investigated for their impact on the activity of azoles by genetic manipulation. Interestingly, a mutation at the same codon (M217I) was previously reported from a Danish CF patient exposed to itraconazole [66]. Another multicenter collection of Aspergillus section Terrei isolates from France also identified azole resistance [67]. None had mutations in the CYP51A gene. Finally, the two azole‐resistant strains reported from the French West Indies were WT for the CYP51A gene [74]. Taken together, these findings suggest that azole resistance in Aspergillus section Terrei is mediated by both CYP51A‐dependent and ‐independent mechanisms. Interestingly, although an environmental route of resistance has not been confirmed for this species, one study showed that azole‐resistant A. terreus clinical isolates with individual Cyp51A mutations (G51A, M217I, or Y491H) also display cross‐resistance with triazole fungicides [110].

3.4.2.2. Amphotericin B

The innate reduced susceptibility of A. terreus to amphotericin B is considered a form of tolerance, as it is not genetically transferable [79]. The drug's action involves interactions with fungal ergosterol and reactive oxygen species, targeting mitochondria. Resistance in A. terreus appears linked to elevated superoxide dismutase activity and enhanced oxidative stress responses [79, 111]. The population of susceptible isolates probably harbours isolates with a tolerant phenotype. Tolerance is not detected by antifungal susceptibility testing, meaning that among the strains with MICs lower than the clinical breakpoint, there are certainly some strains that may have a tolerant phenotype [112].

3.4.2.3. Echinocandin

To our knowledge, there has been no report about the mechanisms of echinocandin resistance in clinical isolates of A. terreus .

3.4.3. Aspergillus Section Nigri

3.4.3.1. Azoles

Only little is known regarding the mechanisms beyond azole resistance in members of the Aspergillus section Nigri. Using gene deletion experiments with a CRISPR‐Cas9 technique, Perez‐Cantero et al. demonstrated that the single deletion of either CYP51A or CYP51B in A. niger decreased the susceptibility to voriconazole by 2 to 16‐fold [113] suggesting that both paralogs are targeted by azoles. As expected from related Aspergillus species, the authors also showed that CYP51A was more involved in the transcriptional response to azole stress than CYP51B. The first study reporting CYP51A mutations in Aspergillus section Nigri dates back to 2008 [22]. Comparison of Cyp51A sequences of azole‐resistant and azole‐susceptible strains of A. awamori revealed amino acid substitutions that seemed unique to azole‐resistant strains (K97T, G427S, and N512K). A similar approach was performed with A. tubingensis and highlighted other mutations (A9V, T321A, P413L, I503V, and L511S). Since then, a handful of other studies have also intended to correlate variations in azole MICs to amino acid polymorphisms in Cyp51A on large sets of clinical isolates belonging to this section [86, 87]. Although some of these substitutions have been only found in azole‐resistant isolates, none have been formally demonstrated to confer azole resistance. On the other hand, partial deletions in CYP51A have been proposed to explain the higher susceptibility of some A. welwitschiae isolates towards azoles [86].

Eventually, besides amino acid polymorphisms in Cyp51A, some studies identified CYP51A overexpression and active efflux through MDR1 and MFS genes as potential drivers of azole resistance in Aspergillus section Nigri [86, 87]. However, despite these encouraging data, the genetic regulation of these transporters is still completely unknown.

3.4.3.2. Amphotericin B and Echinocandin

Mechanisms of amphotericin B or echinocandin resistance have not been explored so far.

3.4.4. Aspergillus Section Nidulantes

3.4.4.1. Azole Antifungals

Aspergillus nidulans, which is closely related to other pathogenic species like A. fumigatus , has been historically used as a model fungus to explore antifungal resistance [114]. One early study, published 20 years ago, used genetic engineering to demonstrate that amplification of the 14 α demethylase gene (referred as pdmA in this study) can confer itraconazole resistance in vitro. Compared to wild‐type controls, itraconazole MICs increased 36‐fold in strains with extra copies of this gene [115]. Yet, these observations remain to be confirmed in clinical isolates. Similarly, early reports have shown that overexpression of Atr transporters (namely AtrA and AtrB) could be responsible for resistance to azole compounds [116, 117]—among other compounds toxic for the fungal cells—in A. nidulans [118]. Although these findings have been observed with azole fungicides, given the close relationship between these azole fungicides and their medical relatives, it is likely that these efflux transporters also contribute to azole resistance in clinical settings.

3.4.4.2. Amphotericin B and Echinocandins

There is currently no report about exploring amphotericin B or echinocandin mechanisms of resistance in A. nidulans .

3.4.5. Other Species

There are yet a few studies exploring the molecular basis of azole resistance in rare Aspergillus species. However, the natural variation M220V in Cyp51A has been proposed to explain the intrinsic azole resistance of A. calidoustus, a member of the section Usti [119].

3.5. Unresolved Issues

3.5.1. Hypothesis About Resistance Selection

Antifungals in human medicine and fungicides in agriculture play a crucial role in the prevention and treatment of mycoses. However, their use is associated with resistance selection. Two selection pathways have been observed in A. fumigatus with azoles [120]. One is the patient route, which consists of selection for resistance in the patients during long‐term use of antifungal drugs. On the other hand, the environmental route relies on the selection of resistant strains outside the human hosts, as typically occurring in the fields with the use of demethylase inhibitor fungicides. Isolates that acquire resistance to fungicides usually display cross resistance to medical antifungals and could be implicated in human infections [121, 122]. There is increasing evidence that the same selection pathways probably exist in non‐fumigatus Aspergillus species.

3.5.1.1. Aspergillus flavus

Previous voriconazole exposure has been reported in patients in whom strains of A. flavus resistant to voriconazole have been isolated. For example, Liu et al. reported the isolation of a voriconazole‐resistant strain after two months treatment by voriconazole in a patient with invasive aspergillosis [51]. Paul et al. reported two isolates from the same patient, one resistant to voriconazole and itraconazole and one resistant to voriconazole only. Two years before, this patient received voriconazole for 4 months followed by itraconazole for more than 6 months for allergic bronchopulmonary aspergillosis [44]. Djenontin et al. reported on both voriconazole‐resistant and itraconazole‐resistant isolates collected in patients with previous azole exposure [56, 57]. Buil et al. reported a patient with aspergilloma who was initially treated with voriconazole for 6 weeks, which allowed the in vivo selection of isogenic azole‐resistant A. flavus isolates [38]. All these arguments confirm the selection of resistance by pressure of medical antifungals in A. flavus .

Finally, as some authors have shown, directed evolution experiments with azoles also support the ability of azoles to select for antifungal resistant isolates [50, 105]. Krishnan et al. performed the selection by culturing 1 x 10⁶ conidia of a susceptible strain for four to 6 days on SDA containing 0.5 mg L⁻¹ voriconazole, followed by screening with RPMI liquid medium containing 4 mg L⁻¹. Voriconazole‐resistant isolates with varying degrees of cross‐resistance to ravuconazole, itraconazole, and posaconazole were noted [50]. Ukai et al. used a different protocol relying on five passage cultures of 7 days each on RPMI agar with 2‐fold serial dilution voriconazole or itraconazole (0.125–16 mg L⁻¹). Isolates harvested from the fifth voriconazole passage showed voriconazole MIC at 16 mg L⁻¹, although no itraconazole resistance could be obtained [105]. Similarly, in a still unpublished work, we showed that resistance selection is possible with both voriconazole and isavuconazole, and to a lesser extent with itraconazole, whereas exposure to posaconazole does not lead to increased MICs despite five culture passages of 7 days.

In addition to the observation of azole‐resistant isolates in patients with previous exposure, it should be noted that resistant strains have also been seldom reported in patients having no history of azole therapy. Indeed, Choi et al. reported seven strains resistant to voriconazole without previous antifungal exposure in the seven patients with otitis media or chronic obstructive pulmonary disease (Table 1) [39]. This last observation supports the hypothesis of an environmental route of resistance selection, as previously reported in A. fumigatus . In addition, several studies have reported environmental strains with high MICs for medical antifungals. As mentioned in the epidemiology chapter, Duong et al., Hermida et al., and Denardi et al. reported environmental isolates with resistance to medical antifungals [37, 40, 42]. Moreover, Meireles et al. showed that A. flavus , exposure to a fungicide pool (thiabendazole + tebuconazole + metconazole), brings to increasing MICs up to 256 times higher for posaconazole and up to 32 times higher for voriconazole even if changes may be reversible after washout. Finally, we have shown that clinical strains resistant to voriconazole have high MICs against the main triazole fungicides used in agriculture (unpublished observations). This cross‐resistance between agricultural fungicides and antifungals implies that resistance selected in agricultural environments has the potential to affect the in vitro susceptibility of A. flavus to medical antifungals.

3.5.1.2. Aspergillus terreus

Azole‐resistant Aspergillus terreus isolates have been reported from a long‐term azole‐treated CF patient, suggesting the possibility of the emergence of azole resistance in vivo, in the patient [66]. On the other hand, azole‐resistant isolates have also been described in environmental samples [68, 79]. In this respect, several demethylase inhibitor fungicides have been tested against WT isolates and isolates resistant to medical triazoles [110]. While WT isolates were susceptible to epoxiconazole, propiconazole, tebuconazole, and difenoconazole, azole‐resistant variants with Cyp51A substitutions (G51A, M217I, and Y491H) displayed increased MICs for these fungicides. These data suggested that epoxiconazole, propiconazole, tebuconazole, and difenoconazole may exert selection pressure for resistance in the environment [110].

3.5.2. Growth and Virulence of Resistant Strains

Resistant strains of micro‐organisms are often less virulent than susceptible ones. In A. fumigatus , however, several experimental studies have reported no fitness cost [123, 124, 125]. This is highlighted by the involvement of azole‐resistant A. fumigatus strains in invasive aspergillosis cases and the high mortality associated with it [126]. Exceptionally, Arendrup et al. reported more virulence in an azole‐susceptible strain than in an isogenic resistance strain [127]. Yet, there is limited data available regarding the fitness cost associated with antifungal resistance in non‐fumigatus Aspergillus species. In a previous publication, we presented the comparison of in vitro growth in liquid RPMI medium of susceptible and resistant A. flavus clinical strains and compared their virulence in the G. mellonella larval model. In vitro growth and virulence of the susceptible strains were higher than those of the resistant clinical strains, supporting a fitness cost in A. flavus [57]. Other phenotypic changes following in vitro exposure to fungicides have been reported by others. For example, Meireles et al. reported that green pigmented colonies of susceptible parent strains changed to white in resistant daughter strains [128]. In A. parasiticus , a common mycotoxin‐producing phytopathogenic fungus, resistant strains to flusilazole (agricultural fungicide) showed changes in mycelial growth, sporulation, and germination compared to susceptible strains [129].

3.5.3. In Vivo and Clinical Implications of In Vitro Resistance – Breakpoints

In the last years and in an era of increased resistance, antifungal susceptibility testing has gained interest in clinical laboratories, allowing clinicians to guide treatments and improve outcomes by facilitating the screening of resistance [130]. Antifungal susceptibility is, for instance, currently recommended in ESCMID guidelines for patients receiving antifungals for proven/probable invasive aspergillosis [131].

Currently available breakpoints established by the EUCAST for systemic antifungals are summarised in Table 4. Note that these breakpoints are not applicable to other methods, such as gradient test strips (the latter method aligning with the CLSI).

TABLE 4.

Overview of antifungal ECOFFs and clinical breakpoints for Aspergillus species according to EUCAST (http://www.eucast.org).

Antifungal	Species	ECOFF (mg L−1)	Clinical breakpoint (mg L−1)
		WT† ≤	S‡≤	R§ >
Amphotericin B	A. fumigatus	1	1	1
	A. flavus	4
	A. nidulans	4
	A. terreus	8
	A. niger	0.5	1	1
Itraconazole	A. fumigatus	1	1	1
	A. flavus	1	1	1
	A. nidulans	1	1	1
	A. terreus	0.5	1	1
	A. niger	2
Voriconazole	A. fumigatus	1	1	1
	A. flavus	2
	A. nidulans	1	1	1
	A. terreus	2
	A. niger	2
Posaconazole	A. fumigatus	0.25	0.125	0.25
	A. flavus	0.5
	A. nidulans	0.5
	A. terreus	0.25	0.125	0.25
	A. niger	0.5
Isavuconazole	A. fumigatus	2	1	2
	A. flavus	2	1	2
	A. nidulans	0.25	0.25	0.25
	A. terreus	1	1	1
	A. niger	4

Abbreviations: R, resistant; S, susceptible; WT, wild type.

3.6. Alternative Therapeutics Against Resistant Strains

With increasing reports of resistant isolates in non‐fumigatus Aspergillus species, alternative treatment strategies are of paramount importance. The two main approaches to address resistance include combination therapy and the use of alternate or new antifungal agents [131, 132]. Combination therapy involves the use of two (or even three) antifungal drugs to exploit potential synergistic effects and enhance efficacy against resistant strains. Meanwhile, the pipeline for new antifungal agents aims to introduce novel compounds with unique mechanisms of action that can effectively target resistant Aspergillus species [133]. These strategies offer promising alternatives in the management of infections caused by resistant non‐fumigatus Aspergillus species.

3.6.1. Treatment of Aspergillus Species With Intrinsic Resistance and Cryptic Species

The treatment of invasive aspergillosis caused by non‐fumigatus Aspergillus species requires careful consideration of intrinsic resistance patterns. Based on the last international recommendations, the following strategies are supported [131]. For Aspergillus terreus, voriconazole, isavuconazole, posaconazole, or itraconazole are recommended, and amphotericin B should be avoided due to intrinsic resistance. Aspergillus calidoustus should be treated with lipid formulations of amphotericin B, avoiding azoles. Aspergillus tubingensis ( A. niger complex) and A. lentulus ( A. fumigatus complex) should not be treated with azole monotherapy due to high MICs for these antifungals. Aspergillus alliaceus ( A. flavus complex) should avoid amphotericin B monotherapy, while the A. niger complex should avoid itraconazole and isavuconazole. Finally, Aspergillus nidulans should be treated with voriconazole due to poor response to amphotericin B. The variable susceptibility patterns between species highlight the fact that accurate species identification and susceptibility testing are crucial to guide effective treatment.

3.6.2. Combination Therapy

There is currently a lack of data on the effectiveness of combination therapies, both in vitro and in vivo, specifically against azole‐resistant non‐fumigatus Aspergillus species. This gap in research poses challenges for developing targeted treatment strategies for infections caused by these resistant strains. However, valuable insights can still be inferred by examining combination therapy results from studies on azole‐resistant A. fumigatus and wild‐type (WT) non‐fumigatus Aspergillus species.

For A. fumigatus , certain combination therapies have shown promise in overcoming resistance, particularly the combination of azole with echinocandins when the level of azole‐resistance is moderate [131, 132], suggesting that similar approaches might be effective against non‐fumigatus species. Additionally, studies on WT non‐fumigatus Aspergillus provide a baseline understanding of how these species might respond to combination treatments. Indeed, several studies have reported synergistic or no interactions for combinations of an azole with an echinocandin against several non‐fumigatus Aspergillus species [134, 135, 136, 137, 138, 139]. Antagonism was rarely observed.

Although combination antifungal therapy can enhance efficacy and broaden the spectrum of activity against resistant strains, it is not always favourable and may sometimes lead to antagonistic interactions. A notable example is the combination of the novel orotomide olorofim with azoles. A recent study [140] demonstrated that the use of azoles in combination with olorofim against A. fumigatus resulted in antagonistic effects in vitro. This antagonism was attributed to the azole‐induced upregulation of the pyrimidine biosynthesis pathway, the target of olorofim, thereby reducing its efficacy. These data highlight the complexity of fungal resistance mechanisms and the need for careful consideration when combination therapies are used to avoid potential antagonistic interactions that could undermine treatment efficacy compared to a recommended monotherapy.

3.6.3. New Antifungals

Several new antifungal drugs are currently in the late stages of development and may offer promising options for managing challenging Aspergillus infections.

Among them, olorofim, a novel antifungal in the orotomide class, has demonstrated strong activity against A. fumigatus , particularly azole‐resistant strains. Additionally, olorofim has shown good efficacy against several non‐fumigatus Aspergillus species belonging to several sections [92, 141, 142, 143]. Nevertheless, higher MICs have been reported in some species such as A. montevidensis and A. chevalieri [141].

While there is no direct data on its activity against non‐fumigatus Aspergillus strains with acquired azole resistance, its efficacy against azole‐resistant A. fumigatus suggests that it could potentially be effective in treating azole‐resistant infections caused by other Aspergillus species as well. Further research is needed to confirm these possibilities.

Similarly, manogepix (formerly APX001), a novel antifungal drug that inhibits the Gwt1 enzyme, has shown promising activity against a wide range of fungal pathogens, including Aspergillus species. It has demonstrated potent in vitro and in vivo activity against A. fumigatus , including azole‐resistant strains [144]. Manogepix also shows good activity against non‐fumigatus Aspergillus species [145, 146]. However, while it is a promising candidate for treating infections caused by Aspergillus species, there is currently no specific data on its efficacy against non‐fumigatus Aspergillus strains with acquired azole resistance.

4. Conclusions

The resistance mechanisms in non‐fumigatus Aspergillus species are far less understood compared to A. fumigatus , although resistance in these species, especially to azoles, is increasingly described. The molecular basis of resistance in these fungi remains poorly characterised, creating numerous gaps in knowledge. Additionally, several key issues, such as the factors driving resistance selection and the clinical relevance of in vitro findings, remain unresolved. Further research is urgently needed to improve our understanding of resistance in these species, which will ultimately lead to better patient management and more effective treatments.

Author Contributions

Elie Djenontin: conceptualization, investigation, writing – original draft, methodology, writing – review and editing, formal analysis. Rose‐Anne Lavergne: conceptualization, writing – original draft, writing – review and editing. Florent Morio: conceptualization, writing – original draft, writing – review and editing. Eric Dannaoui: conceptualization, writing – original draft, writing – review and editing, supervision.

Supporting information

Data S1.

Data Availability Statement

Data sharing is not applicable to this article as no new data were created or analyzed in this study.