Appraisal of Fungus-Derived Xanthoquinodins as Broad-Spectrum Anti-Infectives Targeting Phylogenetically Diverse Human Pathogens

Abstract

Xanthoquinodins make up a distinctive class of xanthone-anthraquinone heterodimers reported as secondary metabolites from several fungal species. Through a collaborative multi-institutional screening program, a fungal extract prepared from a Trichocladium sp. was identified that exhibited strong inhibitory effects against several human pathogens (Mycoplasma genitalium, Plasmodium falciparum, Cryptosporidium parvum, and Trichomonas vaginalis). This report focuses on one of the unique samples that exhibited a desirable combination of biological effects: namely, it inhibited all four test pathogens and demonstrated low levels of toxicity toward HepG2 (human liver) cells. Fractionation and purification of the bioactive components and their congeners led to the identification of six new compounds [xanthoquinodins NPDG A1-A5 (1–5) and B1 (6)] as well as several previously reported natural products (7–14). The chemical structures of 1–14 were determined based on interpretation of their 1D and 2D NMR, HRESIMS, and electronic circular dichroism (ECD) data. Biological testing of the purified metabolites revealed that they possessed widely varying levels of inhibitory activity against a panel of human pathogens. Xanthoquinodins A1 (7) and A2 (8) exhibited the most promising broad-spectrum inhibitory effects against M. genitalium (EC₅₀ values: 0.13 and 0.12 μM, respectively), C. parvum (EC₅₀ values: 5.2 and 3.5 μM, respectively), T. vaginalis (EC₅₀ values: 3.9 and 6.8 μM, respectively), and P. falciparum (EC₅₀ values: 0.29 and 0.50 μM, respectively) with no cytotoxicity detected at the highest concentration tested (HepG2 EC₅₀ > 25 μM).

Graphical Abstract

Broad-spectrum anti-infectives serve important roles for the treatment of a range of human and animal pathogens including bacteria, fungi, viruses, and parasites. Selected examples of compounds that exhibit cross-phylum/multi-kingdom activity against phylogenetically diverse pathogens include nitazoxanide (thiazolide class used to treat Apicomplexa and Metamonada pathogens),¹ paromomycin (amino cyclitol glycoside used to treat Amoebozoa, Euglenozoa, Metamonada, and Platyhelminthes pathogens),² and tinidazole (nitroimidazole class used to treat Amoebozoa, Campylobacterota, and Metamonada pathogens).³ These and other emerging agents^4–7 illustrate the scope of opportunities that exist pertaining to the identification, development, and employment of broad-spectrum anti-infectives.⁸

Similar to the examples noted above, the fungus-derived xanthoquinodins are an intriguing but scantily explored group of natural products that inhibit multiple disease-causing organisms. These heterodimeric metabolites are pragmatically recognized for a distinctive merging of xanthone and anthraquinone moieties.^9,10 The original natural products that helped define this structural archetype were described three decades prior consisting of xanthoquinodins A1 (7), A2 (8), A3 (9), B1 (10), and B2 (11).^9,10 From a broadly inclusive, structure-based perspective, the xanthoquinodins encompass several related named compound series including the acremonidins^11,12 and acremoxanthones,^12–16 beticolins,^17–20 cebetins,^21,22 and engyodontochones.²³

Investigations of the biological effects of xanthoquinodins have revealed that some compounds in this metabolite family are potent inhibitors of the poultry parasite Eimeria tenella,^24,25 while others possess cytotoxic, antimicrobial, and antifungal activities.^26–29 More recently, members from our team determined that compound 7 decreased the growth of the human sexually transmitted parasite Trichomonas vaginalis.³⁰ Considering the diverse structural features embodied within this fungal metabolite family as well as their tentatively useful biological activities, the xanthoquinodins represent a promising lead for the development of a new class of broad-spectrum anti-infectives.

For the purpose of detecting natural product scaffolds with broad-spectrum anti-infective properties, we convened a team of research groups specializing in drug development strategies encompassing four infectious disease-causing organisms: Cryptosporidium parvum, Mycoplasma genitalium, Plasmodium falciparum, and Trichomonas vaginalis. As a key component of our collective efforts, we began by testing a subset of unique fungal natural products (extracts and pure compounds) created largely through the University of Oklahoma, Citizen Science Soil Collection Program,³¹ which is currently composed of >88,000 extracts. Whereas the screening process is still ongoing, these efforts have yielded several promising leads including confirmation that molecules based on the xanthoquinodin scaffold show promise as inhibitors of the four human pathogens targeted by our consortium. Herein we discuss the identification and structure elucidation of previously unreported xanthoquinodins, describe the scope of their inhibitory potential, as well as address the structural features that seemingly influence their broad-spectrum anti-infective effects.

RESULTS AND DISCUSSION

An initial round of in vitro screening focused on testing 1,000 fungus-derived natural product extracts from the University of Oklahoma, Citizen Science Soil Collection. Those tests were performed in parallel against the human parasites M. genitalium and C. parvum. This yielded 22 extracts that met our threshold for inhibition against both target organisms (>60% growth inhibition at 20 μg/mL). These bioactive samples were then screened in a third assay, leading to the identification of an extract derived from a Trichocladium sp. that also showed inhibition of the malaria parasite P. falciparum. This extract was subjected to LC-MS-bioactivity-guided dereplication, revealing several putative xanthoquinodin metabolites. Our team had previously identified xanthoquinodins, such as compound 7, as inhibitors of the sexually transmitted human parasite T. vaginalis.³⁰ In this study, we sought to establish the identities of these new xanthoquinodins and other coeluting metabolites in the bioactive sample. Those efforts led to the purification of six new xanthoquinodin analogues (1–6), along with eight known metabolites (7–14) from the Trichocladium sp. isolate.

Metabolite 1 was obtained as a yellow amorphous solid, and it was determined to have the molecular formula C₃₁H₂₄O₁₂ based on interpretation of its HRESIMS data. Analysis of the HSQC and ¹³C NMR spectra supported the idea that 1 contained 31 unique carbon atoms (Tables 1 and 2). Interpretation of the 2D NMR data (HSQC, HMBC, and COSY) indicated that the structure of compound 1 was similar to xanthoquinodins A1 (7) and A2 (8)⁹ with the major exceptions being the addition of a carbon bearing a hydroxy group [δ_C 78.2 (C-9′)] and the concurrent loss of a nonprotonated (C-9′) carbon. These changes were tied to modest deshielding effects on C-8′ (δ_C 196.6) and C-10′ (δ_C 200.9) in 1 compared with the corresponding carbons in 7 (δ_C 182.9 and 188.9 for C-8′ and C-10′, respectively). Relatedly, both the β-keto–enol tautomeric system (C-8′−10′) and the intramolecularly hydrogen-bonded hydroxy proton (8′-OH: δ_H 14.80, br s) found in 7 were not observed. When CDCl₃ was used as a solvent, we were unable to detect 9′-OH in the ¹H NMR spectrum of 1. Switching to DMSO-d₆, we were able to observe the 9′-OH proton signal (Table S2), which exhibited HMBC correlations to C-8′, −9′, and −14′, in support of the proposed structure for 1 (Figure 1). Comparisons of the chemical shift values for carbons C-2–8 and C-15 of 1 with data collected for xanthoquinodins A1 (7) and A2 (8) indicated that the structure and relative configurations of C-2 and C-3 were unchanged.⁹ This was further supported by the coupling constants recorded for H-3 with H-4_eq (J = 2.0) and H-4_ax (J = 4.0), which corresponded with the values reported for 7. To determine the relative configurations of C-9′, C-11′, and C-14′, a ROESY experiment was employed, providing correlations between 9-OH′, H-12′, and H-13′. This indicated that the hydroxy at C-9′ and the double bond (Δ^12′,13′) occupied the same face of the compound (Figures S1–13).

Table 1.

¹H NMR Spectroscopic Data (500 MHz, CDCl₃) for Compounds 1–6

	1a	2a	3a	4a	5b	6a
3	4.27, dd (4.0, 2.0)	4.55, dd (5.0, 2.0)	4.03, d (10.0)	4.02, d (10.0)	4.93, dd (8.5, 5.5)	4.41, dd (12.5, 5.0)
4	2.13, m	2.33, m	1.85, m	1.85, m	2.33, m	2.30, m
	1.93, m	1.88, m	1.72, m	1.71, m		2.13, m
5	2.82, ddd (19.0, 12.0, 7.0)	3.40, m	2.60, m	2.60, m	2.51, m	2.71, m
		2.41, m
	2.40, dd (19.0, 7.0)
7	-	-	3.13, s	3.13, d (7.5)	3.49, d (17.5)	-
					2.95, d (17.5)
11	-	-	-	-	-	6.12, s
13	6.07, s	6.11, s	6.12, s	6.10, s	6.35, s	-
16	3.69, s	3.68, s	3.74, s	3.73, s	3.64, s	3.74, s
3′	7.53, br s	7.54, br s	7.59, br s	7.54, d (1.5)	7.38 d (2.0)	7.53, br s
5′	7.10, br s	7.11, br s	7.10, br s	7.10, br s	7.15 d (2.0)	7.08, br s
11′	4.93, d (7.5)	4.96, d (7.5)	4.79, d (7.5)	4.92, d (7.5)	4.59 d (7.0)	5.21, d (7.5)
12′	6.42, dd (8.5, 7.5)	6.45, dd (8.5, 7.5)	6.50, dd (8.5, 7.5)	6.43, dd (8.5, 7.5)	6.45, dd (8.5, 7.5)	6.23, dd (8.5, 7.5)
13′	7.01, d (8.5)	7.02, d (8.5)	6.69, d (8.5)	7.02, dd (8.5, 1.0)	6.81, d (8.5)	7.05, d (8.5)
15′	3.18, s	3.20, s	3.07, d (18.0)	3.19, s	3.12, s	3.20, m
			2.91, d (18.0)
16′	2.48, s	2.49, s	2.47, s	2.48, s	2.49, s	2.46, s
6-OH	13.88, s	-	-	-	-	13.94, s
10-OH	12.13, s	12.18, s	12.15, s	12.29, s	12.15, s	11.17 s
6′-OH	10.68, s	10.69, s	11.71, s	10.66, s	10.51, s	10.62, s
9′-OH	nd	nd	nd	nd	8.00, s	nd

^aCDCl₃.

^bDMSO-d₆.

Table 2.

¹³C NMR Spectroscopic Data (100 MHz) for Compounds 1–6

	1a	2a	3a	4a	5b	6a
2	83.9 C	86.3 C	86.5 C	86.5 C	84.6 C	85.7 C
3	66.8 CH	71.0 CH	73.6 CH	73.6 CH	80.9 CH	71.4 CH
4	23.0 CH2	26.3 CH2	25.3 CH2	25.4 CH2	21.9 CH2	23.8 CH2
5	24.4 CH2	31.4 CH2	29.9 CH2	29.7 CH2	27.7 CH2	27.8 CH2
6	180.1 C	198.3 C	176.9 C	175.8 C	176.6 C	179.1 C
7	100.0 C	73.4 C	38.3 CH2	38.4 CH2	39.7 CH2	101.3 C
8	186.5 C	189.1 C	195.2 C	193.0 C	195.2 C	186.5 C
9	105.7 C	105.1 C	105.6 C	106.2 C	106.1 C	106.4 C
10	159.7 C	161.0 C	158.5 C	159.5 C	158.2 C	160.5 C
11	112.9 C	113.6 C	115.8 C	112.2 C	112.4 C	112.3 CH
12	144.0 C	144.8 C	147.7 C	145.1 C	146.4 C	144.3 C
13	109.2 CH	108.6 CH	110.1 CH	110.0 CH	109.6 CH	110.5 C
14	156.3 C	155.6 C	157.4 C	157.6 C	157.7 C	156.3 C
15	170.8 C	168.0 C	170.0 C	169.8 C	169.3 C	169.8 C
16	53.5 CH3	54.0 CH3	53.5 CH3	53.5 CH3	53.9 CH3	53.3 CH3
1′	192.8 C	192.7 C	195.5 C	192.6 C	194.2 C	192.7 C
2′	132.4 C	132.3 C	132.2 C	132.4 C	133.2 C	132.4 C
3′	121.1 CH	121.2 CH	121.0 CH	121.2 CH	120.1 CH	121.3 CH
4′	149.3 C	149.4 C	147.6 C	149.4 C	148.0 C	149.5 C
5′	124.1 CH	124.2 CH	124.4 CH	124.3 CH	124.0 CH	124.2 CH
6′	161.7 C	161.7 C	161.4 C	161.6 C	159.9 C	161.4 C
7′	113.4 C	113.4 C	115.0 C	113.1 C	115.5 C	113.3 C
8′	196.6 C	196.5 C	182.9 C	196.6 C	196.1 C	196.5 C
9′	78.2 C	78.2 C	106.5 C	78.2 C	79.1 C	78.5 C
10′	200.9 C	200.7 C	189.9 C	200.9 C	200.2 C	203.8 C
11′	44.3 CH	44.3 CH	37.8 CH	44.3 CH	44.7 CH	45.2 CH
12′	128.8 CH	129.0 CH	131.4 CH	128.8 CH	128.9 CH	129.0 CH
13′	131.0 CH	130.9 CH	132.7 CH	131.0 CH	131.1 CH	130.7 CH
14′	56.6 C	56.5 C	49.9 C	56.4 C	57.3 C	56.6 C
15′	36.8 CH2	36.9 CH2	39.0 CH2	36.9 CH2	36.0 CH2	36.8 CH2
16′	22.3 CH3	22.3 CH3	22.1 CH3	22.3 CH3	21.7 CH3	22.3 CH3

^aCDCl₃.

^bDMSO-d₆.

Figure 1.

Key HMBC and COSY correlations of compounds 1–6.

The absolute configuration of 1 was investigated by comparisons of its electronic circular dichroism (ECD) spectrum with calculated values for a range of chemical species by using quantum mechanical time-dependent density functional theory (TDDFT) (Figure 2A). To this end, ECD calculation experiments were carried out on several putative stereoisomers and partial structures (1a–1e, Supporting Information Figures S7-1 and 12a–d, Supporting Information Figure S7-2) to better understand how the various chromophoric features in 1 influenced its ECD spectrum. Results from these efforts indicated the stereochemical relationship of the C-9′ hydroxy group and double bond (Δ12′,13′) was crucial for producing the negative Cotton effect between 300 and 350 nm. Further analysis showed that an S configuration at C-3 helped deliver a positive Cotton effect between 250 and 300 nm, which appeared unaffected by the configuration of the C-2 methyl ester group. This configurational assignment aligned well with previous reports that most xanthoquinodins bearing a C-3 hydroxy group possessed 3S configurations.^9,24–28 Based on these observations, the absolute configuration for 1 was established as 2R,3S,9′R,11′S,14′R, and the compound was assigned the trivial name xanthoquinodin NPDG A1 (1).

Figure 2.

(A–D) Experimental ECD spectrum of compounds 1–9 and calculated ECD spectrum of 1, 2, 5, and chaetomanone.

Compound 2 was obtained as a yellow amorphous solid, and its molecular formula was determined to be C₃₁H₂₄O₁₃ based on analysis of its HRESIMS data. An investigation of its 1D (Tables 1 and 2) and 2D NMR (Figure 1) data revealed that the structure of 2 was similar to that of 1; however, compound 2 possessed an additional hydroxy group hypothesized to be located on C-7. To confirm this, the HMBC experiment was performed a second time using DMSO-d₆ as the solvent, which enabled the detection of key correlations from 7-OH to C-2, C-7, and C-8 (Table S2). The relative configuration of the hydroxy group was determined based on analysis of ROESY data, which included correlations among protons H-3/7-OH/H-5_eq/H-4_ax/Me-16, indicating that 7-OH and Me-16 occupied the same face of the compound (Figure 3). The relative configurations of C-3 and C-4 were determined by analysis of coupling constant data, which showed that H-3 coupled with H-4_eq (J = 2.0) and H-4_ax (J = 5.0) in a manner consistent with compounds 1 and 7 (vide supra). ROESY correlations among H-12′, H-13′, and OH-9′ aided in confirming their co-occurrence on the same face of compound 2. Similar to our analysis for compound 1, the absolute configuration of 2 was investigated by a comparative analysis of experimental and calculated ECD data (Figure 2B), which led to assigning the metabolite a 2S,3S,7R,9′R,11′S,14′R configuration. Compound 2 was given the trivial name xanthoquinodin NPDG A2.

Figure 3.

Key ROESY correlations observed for compound 2.

Compound 3 was purified as a yellow amorphous solid, and it was assigned the molecular formula C₃₁H₂₆O₁₂ based on interpretation of its HRESIMS data. Comparisons of its ¹H and ¹³C NMR data (Tables 1 and 2) with those obtained for other xanthoquinodin analogues suggested that compound 3 shared a similar structure with xanthoquinodins A3 (9)⁹ and A5.²⁶ The major differences between compounds 3 and 9 were the changes in chemical shifts attributed to C-2 through C-6, which indicated that the γ-lactone in 9 was opened and replaced by a 1-hydroxybutyric acid moiety. This was further supported by the absence of an HMBC correlation between H-3 and C-6, as had been observed with compound 9.⁹ During the period spent collecting NMR data, a second set of low-intensity peaks emerged in the NMR spectra. Upon recollection and analyses of the proton and carbon chemical shift data, it was found that the new signals precisely correlated with spectra separately collected for compound 9. These observations suggest that 3 can be partially converted into 9 in a solvent (DMSO-d₆) at room temperature (Figures S3–10).The ECD spectrum for 3 exhibited a negative Cotton effect between 200 and 250 nm and a positive Cotton effect centered near 340 nm, which were observed to be similar to the ECD spectra for compounds 7–9 (Figure 2C). Calculated ECD data prepared for 3 and 7–9 showed strong matches to experimental results (Figures S7-3). Thus, the absolute configuration of 3 was determined to be 2R,3S,11′S,14′R, and the compound was given the trivial name xanthoquinodin NPDG A3.

Xanthoquinodin NPDG A4 (4) was obtained as a yellow amorphous solid, and its molecular formula was inferred to be C₃₁H₂₆O₁₃ based on an analysis of its HRESIMS data. Considering the metabolite’s molecular formula, as well as comparing its 1D and 2D NMR data with those obtained for 3, we determined that a key structural difference between these metabolites was the addition of a new alcohol group. Analysis of its NMR data enabled us to posit that the new organic functionality in 4 was appended to C-9′ with the adjacent enol present in 3 having been converted to a ketone in 4. This was supported by a distinctive chemical shift recorded for C-9′ (δ_C = 78.2 ppm), which was consistent with values recorded for the C-9′ positions in analogues 1 and 2 (Table 2).

Similar to the phenomenon we noted while collecting the NMR spectra for compound 3, it was observed that a new set of peaks had emerged in the material used for our analysis of sample 4 (DMSO-d₆). However, distinct from the instance with compound 3, we saw that the new set of “minor” signals continued to increase in intensity, eventually becoming the dominate (“major”) signals, as measured by 1D NMR. This enabled us to proceed with gathering a full 1D and 2D NMR data set for the emergent “major” component. While these events unfolded, serendipity led us to observe that the NMR data being collected for another presumptive xanthoquinodin analogue purified from a separate fraction, compound 5, yielded 1D and 2D NMR spectra (acquired in DMSO-d₆) matching the data being collected for the new “major” product originating from sample 4. The data we obtained for these two compounds were similar but not identical to values reported for chaetomanone (C-7′: 115.5 vs 118.0, C-9′: 79.1 vs 81.5, C-10′: 200.2 vs 202.9, C-13′: 131.1 vs 135.5, C-14′: 57.3 vs 54.4, and C-15′: 36.0 vs 30.6; note that the first value given is for compound 5 and values for chaetomanone are listed second).³² This led us to conjecture that 5 might be a stereoisomer of chaetomanone.

Taking these observations into account, we returned to interpreting the NMR data for compounds 4 and 5 (Tables 1 and 2, Figure 1) and determined that 4 possessed a 1-hydroxybutyric acid (C2–C6) moiety, which had undergone lactonization to form compound 5. Interpretation of the HRESIMS data for 5 supported this assessment, with its molecular formula established to be C₃₁H₂₄O₁₂ (an increase of one unit of unsaturation relative to 4). ROESY correlation data were used to probe the relative configurations of both compounds, with key correlations establishing that H-12′, H-13′, and 9′-OH occupied the same relative faces of the two compounds (Figures S4–12 and S5–9). Analyses of the ECD spectra for 4 and 5 revealed positive Cotton effects between 240 and 300 nm and negative Cotton effects centered near 362 nm (Figure 2D). Considering these results as well as biogenic factors, it is proposed that both 4 and 5 have 2R,3S,9′R,11′S,14′R absolute configurations. The compounds were assigned the trivial names xanthoquinodins NPDG A4 (4) and NPDG A5 (5).

Metabolite 6 was purified as an amorphous yellow solid, and its molecular formula was determined to be C₃₁H₂₄O₁₂ based on interpretation of its HRESIMS data. Examination of the 1D and 2D NMR data for 6 indicated that this compound was structurally similar to xanthoquinodins B1 (10) and B2 (11)⁹ with the major difference being the addition of a hydroxy group at C-9′ (Tables 1 and 2). HMBC correlations from H-11′ to C-13 and C-14 established the presence of a bond between C-11′ and C-13 as noted among other members of the xanthoquinodin “B” series (Figure 1). The relative configuration of 6 in the xanthone portion of the compound was initially inferred based on the strong similarities of the carbon chemical shift values for C-2 through C-7 and C-15 with those observed for xanthoquinodin B2 (11). This proposal was supported in part by proton coupling data showing that H-3 coupled with H-4_eq (J = 5.0) and H-4_ax (J = 12.5), which agreed with data recorded for compounds 8 and 11. Analysis of the ECD spectrum for 6 revealed a pattern of Cotton effects similar to that of 1 (Figure 2A). Thus, the absolute configuration of 6 was determined to be 2S,3S,9′R,11′S,14′R, and it was given the trivial name xanthoquinodin NPDG B1 (6). Notably, compounds 1 and 6 have been reported to be chemically interconvertible as is the case for other members of the xanthoquinodin family.⁹

In addition to compounds 1–6, eight known compounds were identified from this fungus including xanthoquinodin A1 (7),⁹ xanthoquinodin A2 (8),⁹ xanthoquinodin A3 (9),^9,24 xanthoquinodin B1 (10),⁹ xanthoquinodin B2 (11),⁹ gonytolide C (12),³³ and two emodin-bianthrones (13–14).³⁴ The identities of these compounds were determined based on comparisons of their physicochemical and spectroscopic data with the reported values.

Compounds 1–9 and 12–14 were tested for their inhibitory effects against M. genitalium, C. parvum, P. falciparum, and T. vaginalis (Table 3). Human hepatocytes (HepG2 cells) were used to evaluate each compound’s cytotoxicity and calculate the selectivity index (SI) values for each pathogen (Table 3). Several members of the xanthoquinodin series (1–9) exhibited potent anti-infective properties against all of the pathogens tested with EC₅₀ values at, or below, micromolar concentrations. The archetypal xanthoquinodin compounds 7 and 8 showed the greatest potencies against M. genitalium (EC₅₀ values of 0.13 ± 0.02 and 0.12 ± 0.01 μM, respectively), C. parvum (EC₅₀ values of 5.2 ± 1.8 and 3.5 ± 0.5 μM, respectively), P. falciparum (EC₅₀ values of 0.29 ± 0.01 and 0.50 ± 0.03 μM, respectively), and T. vaginalis (EC₅₀ values of 3.9 ± 0.3 and 6.8 ± 1.0 μM, respectively). In general, the newly described analogues (1–6) were found to be less active or inactive at the concentrations tested. The bianthrones (13–14) exhibited modest activity against P. falciparum but were found to be significantly less selective as demonstrated by their cytotoxic effects on HepG2 cells (EC₅₀ values of 13.8 and 10.2 μM, respectively) and lower selectivity index values (SI values of 8.6 and 9.1, respectively). In comparison, no cytotoxicity was seen in HepG2 for the xanthoquinodins, at concentrations extending up to 25 μM.

Table 3.

Bioassay Results (EC₅₀ Values Expressed in μM ± SEM) for the Compounds (1–9 and 12–14) Tested against Parasites (T. vaginalis, M. genitalium, C. parvum, and P. falciparum) and Human Cells (HepG2)^a

	T. vaginalis		M. genitalium		C. parvum		P. falciparum		HepG2
compound	EC50	SIb	EC50	SI	EC50	SI	EC50	SI	EC50
1	>10	ndc	>10	nd	>10	nd	>5	nd	>25
2	>10	nd	>10	nd	>10	nd	>5	nd	>25
3	>10	nd	>10	nd	>10	nd	>5	nd	>25
4	>10	nd	>10	nd	>10	nd	>5	nd	>25
5	>10	nd	>10	nd	>6	nd	>5	nd	>25
6	>10	nd	>10	nd	>10	nd	>5	nd	>25
7	3.9 ± 0.3	>6.4	0.13 ± 0.02	>62.5	5.2 ± 1.8	>4.8	0.29 ± 0.01	>86	>25
8	6.8 ± 1.0	>3.7	0.12 ± 0.01	>125	3.5 ± 0.5	>7.1	0.50 ± 0.03	>50	>25
9	>10	nd	ntd	-	nt	-	>5	nd	>25
12	>10	nd	>10	nd	>40	nd	>5	nd	>25
13	>10	nd	>10	nd	>7.2	nd	1.65 ± 0.14	8.6	13.8
14	>10	nd	>10	nd	>2.4	nd	1.12 ± 0.16	9.1	10.2

^aPositive controls: metronidazole for T. vaginalis (EC₅₀ = 3.5 ± 0.5 μM), azithromycin for M. genitalium (EC₅₀ = 0.0005 ± 0.0001 μM), BKI-1294 for C. parvum (EC₅₀ = 1.1 ± 0.2 μM), and chloroquine for P. falciparum (EC₅₀ = 0.17 μM).

^bSelectivity index value.

^cNot determined.

^dNot tested.

Comparisons of the structures representing the most potent and selective compounds against the less active/inactive substances revealed two major chemical features that appear to strongly influence their biological activities. First, the introduction of an alcohol group to the C-9′ position of the xanthoquinodins with the concomitant loss of the C-8′/C-9′ enol resulted in a profound loss of biological activity. Second, changes to the cyclohex-1-ene-1,4-diol system (C-2 through C-7 ring system in the “xanthone” portion of xanthoquinodins wherein cyclization had not occurred or the atoms had undergone transformation to γ-lactone-containing structures) resulted in significant or complete losses of activity. Together, these results suggest that two enol moieties, one in the “anthraquinone” portion and one in the “xanthone” portion of the xanthoquinodins, engaged in intramolecular H-bonding to adjacent ketones, are essential for their potent inhibitory effects against the pathogens examined in this study.

Comparing the results of our biological studies to data reported for 7–9 and other xanthoquinodins, we found a handful of seeming incongruencies warranting discussion. For example, the mammalian cell cytotoxicity values reported for several xanthoquinodins including 7, 8, 10, and 11 indicate a more concerning potential for toxicity than we observed.^{24–27,29,35,36} It is possible that these dissimilarities may be due to the different cell lines tested or the experimental conditions employed, making further testing a necessity to better understand the safety threshold of this compound class. It is worth noting that in our studies compounds were only tested as high as 25 μM in order to maintain solubility while not exceeding a nontoxic concentration of DMSO, a consideration that may not have been necessary with previous efforts. Additionally, our assertion that dual H-bonded enols are essential for potent biological activities are challenged by reports that xanthoquinodins bearing single enols exhibit substantial inhibitory effects against a variety of organisms.^24–29 The dissimilarities between our observed structure–activity trends and those for other xanthoquinodin series suggest that the inhibition of different pathogens/cell types may involve distinct modes of target engagement. It is anticipated that additional studies aimed at delving deeper into the structure–activity relationship (SAR) of xanthoquinodins will help discern the chemical features that control the biological activities of these compounds.

As a point of concern, we would like to draw attention to some seemingly anomalous NMR chemical shift values and interpretation of 2D NMR correlation data reported for xanthoquinodin B9.²⁸ Specifically, we encountered a number of unusual carbon chemical shift values described for the “anthraquinone” portion of the molecule, which was hypothesized to be a “B” type xanthoquinodin (i.e., it was presumed to share the same chemical scaffold as compounds 10, 11, and xanthoquinodin B3). The incongruous ¹³C NMR data include assignments for C-1′ (Δ ≈ 5 ppm from expected), C-8′ (Δ ≈ 7 ppm from expected), C-10′ (Δ ≈ 11 ppm from expected), C-11′ (Δ ≈ 8 ppm from expected), and C-15′ (Δ ≈ 3 ppm from expected), combined with the presence of unusual four-bond HMBC correlations (H-11′ to C-8′ and H-11 to C-8), suggested to us that this structure should be considered with caution pending further analysis.

In summary, we have found xanthoquinodins to be a promising group of compounds that merit further investigation as possible broad-spectrum anti-infective agents targeting multiple human pathogens. Several outstanding questions remain concerning the relative toxicity and structural features that are essential for their biological effects. A combination of approaches involving synthetic/semisynthetic methods as well as procuring additional analogues from nature are anticipated to help clarify the potential utility of these compounds as therapeutic leads.

EXPERIMENTAL SECTION

General Experimental Procedures.

Optical rotation measurements were made by using a Rudolph Research Autopol III automatic polarimeter. NMR spectra were obtained on Varian NMR spectrometers (400 and 500 MHz for ¹H NMR measurements and 100 and 125 MHz for ¹³C NMR measurements). HRESIMS data were obtained on an Agilent 6538 high-mass-resolution QTOF mass instrument. ECD spectra were obtained on a JASCO J-715 circular dichroism spectrometer. Vacuum column chromatography was performed over a silica gel (VWR, 40–60 μm, 6 Å) and HP20ss gel (Sorbtech). The preparative HPLC system was equipped with Shimadzu SCL-10A VP pumps and a system controller, using a Gemini 5 μm C₁₈ column (210 Å, 250 × 21.2 mm) with a flow rate of 10 mL/min. Semipreparative HPLC separations were performed on a Waters system (1525 binary pumps and Waters 2998 photodiode array detectors) using a Gemini 5 μm C₁₈ (110 Å, 250 × 10 mm) or F5 (110 Å, 250 × 10 mm) column with a flow rate of 4 mL/min. All solvents used were of ACS grade or better.

Fungal Isolate and Fermentation.

The Trichocladium sp. isolate (TN09213 RBM-1) was obtained from a soil sample collected near Lebanon, Tennessee, by a citizen scientist. The fungus was identified by collecting fresh mycelium from a Petri plate and subjecting the sample to homogenization in TE buffer (10 mM EDTA HCl, 0.1 mM EDTA, pH 8.0) with zirconium oxide beads in a Bullet blender (MidSci #BBY24M). Fungal DNA was collected, and the ITS region (i.e., ITS1, 5.8S, and ITS2 regions) was amplified by PCR for sequencing. The resulting sequence data were compared to fungal sequences contained in GenBank, which led to two 100% identity matches to isolates described as Trichocladium asperum and Trichocladium griseum. Sequence data were deposited in GenBank (T. asperum: GenBank accession no. MZ230232).

To prepare the isolate for chemical purification studies, the fungus was recovered from cryogenic storage (stored as mycelium suspended in vials with 20% aqueous glycerol at −80 °C). Following recovery on Czapek agar plates (30 g sucrose, 2 g NaNO₃, 1 g K₂HPO₄, 0.5 g MgSO₄·7H₂O, 0.5 g KCl, 0.01 g FeSO₄·7H₂O, 50 mg chloramphenicol, 1 L DI H₂O), Petri plates containing lawns of fungal mycelium were aseptically cut into small pieces (~1 cm²) to use as inoculum for scale-up cultures. Scale-up cultures were prepared by charging mycobags (Unicorn Bags, Plano, TX, USA) with monolayers of autoclaved Cheerios breakfast cereal, supplemented with a 0.3% sucrose solution and 0.005% chloramphenicol, and then seeding the bags with the cut-up pieces of fungal culture. Each mycobag provides approximately 1320 cm² of surface area to support fungal culture growth. For the experiments reported in this study, three mycobags were prepared, and the fungal samples were grown at room temperature with constant lighting for 4 weeks.

Extraction and Purification of Compounds from T. asperum.

Fungal biomass was extracted with 2 L of EtOAc (×3) at room temperature. The resulting organic soluble materials were pooled before being subjected to evaporation under a vacuum. This yielded 35 g of EtOAc-soluble material (fraction A). Fraction A was subjected to silica gel vacuum column chromatography with elution performed by using dichloromethane (fraction B), dichloromethane-MeOH (10:1) (fraction C), and MeOH (fraction D). Fraction C (4 g) was further separated by HP20ss gel vacuum column chromatography into five samples: fractions E (30% MeOH), F (50% MeOH), G (70% MeOH), H (90% MeOH), and I (100% MeOH). Fraction I (340 mg) was further subjected to preparative HPLC (C₁₈, gradient elution with 80–100% MeOH-H₂O over 15 min using a 10 mL/min flow rate) to afford six subfractions (I1–I6). Subfraction I1 (50 mg) was subjected to semipreparative HPLC [C₁₈, isocratic MeCN-H₂O (50:50), flow rate: 4 mL/min] yielding compounds 4 (2 mg, t_R = 8 min), 12 (2 mg, t_R = 8.5 min), 2 (3 mg, t_R = 9 min), 5 (6 mg, t_R = 15 min), and 1 (6 mg, t_R = 17 min). Compound 3 (2 mg, t_R = 18 min) was purified from subfraction I2 (25 mg) by semipreparative HPLC [C₁₈, isocratic MeCN-H₂O (60:40), flow rate: 4 mL/min]. Compound 6 (6 mg, t_R = 16 min) was purified from subfraction I3 (20 mg) by semipreparative HPLC [C₁₈, isocratic MeCN-H₂O (55:45), flow rate: 4 mL/min]. Subfraction I4 (30 mg) was subjected to semipreparative HPLC [C₁₈, isocratic MeCN-H₂O (65:35), flow rate: 4 mL/min], yielding compounds 13 (2 mg, t_R = 13 min) and 14 (3 mg, t_R = 16 min). Fraction D (4 g) was further subjected to HP20ss gel vacuum column chromatography, which provided five fractions: fractions K (30% MeOH), L (50% MeOH), M (70% MeOH), N (90% MeOH), and O (100% MeOH). Fraction O (700 mg) was subjected to preparative HPLC (C₁₈, gradient elution with 80–100% MeOH-H₂O over 15 min using a 10 mL/min flow rate) to generate six subfractions (O1–O6). Subfraction O4 (55 mg) was subjected to semipreparative HPLC [F5, isocratic MeCN-H₂O (60:40), flow rate: 4 mL/min] to give compounds 9 (1 mg, t_R = 14 min), a mixture of 10 and 11 (1 mg, t_R = 16 min), and 7 (10 mg, t_R = 20 min). Compound 8 (2 mg, t_R = 12 min) was obtained from subfraction O5 (20 mg) by semipreparative HPLC [C₁₈, isocratic MeCN-H₂O (80:20), flow rate: 4 mL/min].

Xanthoquinodin NPDG A1 (1).

An amorphous yellow solid; [α]²⁵_D + 112 (c 0.1, MeOH); ECD (MeOH) λ_max (Δε) 208 (−24.2), 225 (−18.3), 284 (+12.5), 323 (−2.0), 355 (+1.6); ¹H NMR (500 MHz, CDC1₃) and ¹³ C NMR (100 MHz, CDC1₃); see Tables 1 and 2; HRESIMS m/z 589.1351 [M + H]⁺ (calcd for C₃₁H₂₅O₁₂, 589.1341; mass error −1.7 ppm).

Xanthoquinodin NPDG A2 (2).

An amorphous yellow solid; [α]²⁵_D + 32 (c 0.1, MeOH); ECD (MeOH) λ_max (Δε) 208 (−23.5), 224 (−17.0), 283 (+8.4), 363 (−1.8); ¹H NMR (500 MHz, CDC1₃) and ¹³C NMR (100 MHz, CDC1₃), see Tables 1 and 2; HRESIMS m/z 605.1287 [M + H]⁺ (calcd for C₃₁H₂₅O₁₃, 605.1290; mass error +0.5 ppm).

Xanthoquinodin NPDG A3 (3).

An amorphous yellow solid; [α]²⁵_D + 156 (c 0.1, MeOH); ECD (MeOH) λ_max (Δε) 209 (−23.0), 225 (−19.5), 284 (+2.8), 340 (+3.6), 396 (+4.1); ¹H NMR (500 MHz, CDC1₃) and ¹³ C NMR (100 MHz, CDC1₃); see Tables 1 and 2; HRESIMS m/z 591.1498 [M + H]⁺ (calcd for C₃₁H₂₇O₁₂, 591.1497; mass error −0.2 ppm).

Xanthoquinodin NPDG A4 (4).

An amorphous yellow solid; [α]²⁵_D + 64 (c 0.1, MeOH); ECD (MeOH) λ_max (Δε) 207 (−50.6), 225 (−35.7), 282 (+21.2), 359 (−2.4); ¹H NMR (500 MHz, CDC1₃) and ¹³C NMR (100 MHz, CDC1₃); see Tables 1 and 2; HRESIMS m/z 607.1451 [M + H]⁺ (calcd for C₃₁H₂₇O₁₃, 607.1446; mass error −0.8 ppm).

Xanthoquinodin NPDG A5 (5).

An amorphous yellow solid; [α]²⁵_D + 38 (c 0.1, MeOH); ECD (MeOH) λ_max (Δε) 207 (−35.6), 224 (−29.6), 283 (+13.0), 363 (−2.2); ¹H NMR (500 MHz, DMSO-d₆) and ¹³C NMR (100 MHz, DMSO-d₆); see Tables 1 and 2; HRESIMS m/z 611.1170 [M + Na]⁺ (calcd for C₃₁H₂₄NaO₁₂, 611.1160; mass error −1.6 ppm).

Xanthoquinodin NPDG B1 (6).

An amorphous yellow solid; [α]²⁵_D + 80 (c 0.1, MeOH); ECD (MeOH) λ_max (Δε) 203 (−10.4), 231 (−1.4), 279 (+7.1), 325 (−4.3), 358 (+2.9); ¹H NMR (500 MHz, CDC1₃) and ¹³ C NMR (100 MHz, CDC1₃); see Tables 1 and 2; HRESIMS m/z 589.1345 [M + H]⁺ (calcd for C₃₁H₂₅O₁₂, 589.1341; mass error −0.7 ppm).

Trichomonas vaginalis Assay.

Trichomonas vaginalis Donne (PRA-98) from the American Type Culture Collection (ATCC, Bethesda, MD) was grown at 37 °C in filter sterilized Keister’s Modified TYI-S33 medium (20 g of casein, 10 g of yeast, 10 g of glucose, 2 g of NaCl, 0.6 g of KH₂PO₄, 1 g of K₂HPO₄, 2 g of l-cysteine HCl, 0.2 g of l-ascorbic acid, 23 mg of ferric ammonium citrate, 100 mL of heat inactivated fetal bovine serum, 0.52 g of bovine bile salts in 1 L of Millipore water). Micro aerophilic conditions were maintained with the use of BD GasPak EZ Campy sachets. Samples consisting of 4 × 10⁴ trichomonads per well were treated with DMSO, 25 μM metronidazole, or pure compounds, not exceeding 0.5% DMSO. After a 17 h incubation, 100 μL of fixing solution (1% glutaraldehyde, 5 μM propidium iodide, and 5 μM acridine orange in PBS) was applied to each well. Plates of treated samples were placed on an orbital shaker for 30 s to disperse clumps, followed by a 3 h incubation at 37 °C. Cells were subsequently imaged using a PerkinElmer Operetta high-content imaging system, and data were analyzed using the Harmony 3.5.1 software package as previously described.³⁰ EC₅₀ values were calculated by using GraphPad Prism version 4.03.

Mycoplasma genitalium Assay.

Samples were tested for activity against M. genitalium strain G37 using a microbroth dilution assay similar to procedures described by Hamasuna et al.³⁷ Briefly, M. genitalium frozen stocks were inoculated into SP-4 broth³⁸ and incubated for 2 days at 37 °C under an atmosphere of 5% CO₂ at which point the phenol red indicator turned orange indicating log phase growth. Adherent bacteria were scraped, suspended in the culture supernatant, and passed through a 0.45 μm filter to remove the cell aggregates. The inoculum was diluted 1:200 with fresh SP-4 broth containing 7.5% fetal bovine serum, and 100 μL (~10⁴ CFU) was added to wells of a flat bottom 96-well plate containing 100 μL of test agent diluted in the same medium. Plates were incubated for 6–7 days at 37 °C under an atmosphere of 5% CO₂ before being examined for color change. To determine EC₅₀ values, growth inhibition by 2-fold serial dilutions of compounds tested in triplicate was determined by qPCR targeting the 16s rRNA gene³⁹ as compared to untreated wells. Total DNA in each well was isolated using the MasterPure Complete DNA and RNA Purification kit (Lucigen, Middleton, WI), and M. genitalium genomes were quantified in triplicate as compared to a quadruplicate standard curve. Percent inhibition was calculated, and EC₅₀ values were determined with GraphPad Prism 9.3.1 (GraphPad Software, San Diego, CA).

C. parvum Assay.

Inhibition of C. parvum growth and EC₅₀ value determination were performed with a Nanoluciferase-tagged C. parvum parasite in HCT-8 cells. Oocysts were obtained from the University of Arizona after propagation in newborn calves. HCT-8 cells were seeded into a 384-well plate and allowed to grow for 48 h to reach 50–75% confluence. Oocysts were activated by a 15 min incubation in 10% bleach (0.6% sodium hypochlorite) at room temperature and then washed with DPBS. Aliquots consisting of 5 × 10³ oocysts per well were applied to 384-well plates with RPMI-1640 medium supplemented with 10% heat inactivated fetal bovine serum and 1% penicillin-streptomycin solution (10,000 units/mL of penicillin and 10,000 μg/mL of streptomycin) and varying concentrations of test compounds and incubated for 48 h at 37 °C with an atmosphere of 5% CO₂. Luciferase-induced light emission was then determined by lysing the resulting cell monolayers, adding Nano-Glo luciferase reagent (Promega, Madison, WI, USA), and detecting the signal on an EnVision Multilabel Plate Reader (PerkinElmer, Waltham, MA, USA). Controls included infected wells with no compound addition, and wells with no C. parvum propagules for background value establishment. The percentage of growth versus infected cells with no inhibitor was calculated for each concentration of compound, and the half maximal effective concentration (EC₅₀) values were determined via sigmoidal dose response with variable slope using GraphPad Prism version 6.07 (GraphPad Software, La Jolla, California, USA).

Plasmodium falciparum Assay.

Culturing of P. falciparum Dd2 was performed as previously described⁴⁰ with no deviations. Antiplasmodial EC₅₀ results were also determined per previous reference using a SYBR Green I assay performed with asynchronous stage parasites. Fluorescence was measured using a Synergy Neo2 multimode reader (BioTek, Winsooki, VT, USA), with chloroquine-treated and untreated controls. Dose–response plots and EC₅₀ values were generated using CDD Vault (Collaborative Drug Discovery, Burlingame, CA).

Human Cell Cytotoxicity Assay.

Mammalian cell cytotoxicity was assessed in HepG2 human hepatocytes using an MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium) based assay as previously described.⁴⁰ Compounds were tested in triplicate at a maximum concentration of 25 μM to avoid issues of solubility while maintaining a nontoxic concentration of DMSO. Absorbance was determined using a Synergy Neo2 multimode reader (BioTek), with a 5% Triton X-100-lysed zero viability control, and a no-treatment control. Dose–response plots and EC₅₀ values were generated using CDD Vault.

ECD Calculations.

Compound conformational searches were carried out using ComputeVOA (BioTools, Inc.) at the molecular mechanics level (MMFF). Conformers were further optimized at the B3LYP/6–31G (d,p) level with Gaussian 09 (Gaussian Inc.). All stable conformers were subjected to an ECD calculation by time-dependent DFT at the B3LYP/6–311+g(d,p) level. The ECD spectra were added together after a Boltzmann statistical weighting using SpecDis 1.71 (sigma value of 0.3 eV). After a UV-shift correction was applied, the computed ECD spectra were compared to the experimentally derived ECD curves.

ABBREVIATIONS

NMR

nuclear magnetic resonance

ECD

electronic circular dichroism

HRESIMS

high-resolution electrospray ionization mass spectrometry

SI

selectivity index

VLC

vacuum liquid chromatography

ITS

internal transcribed spacer

SAR

structure–activity relationship

HSQC

heteronuclear single quantum coherence spectroscopy

HMBC

heteronuclear multiple bond correlation spectroscopy

COSY

homonuclear correlation spectroscopy

ROESY

rotating frame nuclear Overhauser effect spectroscopy