Inhibiting protein prenylation with benzoxaboroles to target fungal plant pathogens

Article abstract of DOI:10.1021/acschembio.0c00290

Fungal pathogens pose an increasing threat to global food security through devastating effects on staple crops and contamination of food supplies with carcinogenic toxins. Widespread deployment of agricultural fungicides has increased crop yields but is driving increasingly frequent resistance to available agents and creating environmental reservoirs of drug-resistant fungi that can also infect susceptible human populations. To uncover non-cross-resistant modes of antifungal action, we leveraged the unique chemical properties of boron chemistry to synthesize novel 6-thiocarbamate benzoxaboroles with broad spectrum activity against diverse fungal plant pathogens. Through whole genome sequencing of Saccharomyces cerevisiae isolates selected for stable resistance to these compounds, we identified mutations in the protein prenylation-related genes, CDC43 and ERG20. Allele-swapping experiments confirmed that point mutations in CDC43, which encodes an essential catalytic subunit within geranylgeranyl transferase I (GGTase I) complex, were sufficient to confer resistance to the benzoxaboroles. Mutations in ERG20, which encodes an upstream farnesyl pyrophosphate synthase in the geranylgeranylation pathway, also conferred resistance. Consistent with impairment of protein prenylation, the compounds disrupted membrane localization of the classical geranylgeranylation substrate Cdc42. Guided by molecular docking predictions, which favored Cdc43 as the most likely direct target, we overexpressed and purified functional GGTase I complex to demonstrate direct binding of benzoxaboroles to it and concentration-dependent inhibition of its transferase activity. Further development of the boron-containing scaffold described here offers a promising path to the development of GGTase I inhibitors as a mechanistically distinct broad spectrum fungicide class with reduced potential for cross-resistance to antifungals in current use.

Full text of DOI:10.1021/acschembio.0c00290

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Articles
Inhibiting Protein Prenylation with Benzoxaboroles to Target
Fungal Plant Pathogens
Sang Hu Kim, Chunliang Liu, Yasheen Zhou, Yong-Kang Zhang, Cari McGregor, Luke Steere,
Brittany H. Frederick, C. Tony Liu, Luke Whitesell, and Leah E. Cowen*
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ABSTRACT: Fungal pathogens pose an increasing threat to
global food security through devastating effects on staple crops and
contamination of food supplies with carcinogenic toxins. Wide-
spread deployment of agricultural fungicides has increased crop
yields but is driving increasingly frequent resistance to available
agents and creating environmental reservoirs of drug-resistant fungi
that can also infect susceptible human populations. To uncover
non-cross-resistant modes of antifungal action, we leveraged the
unique chemical properties of boron chemistry to synthesize novel
6-thiocarbamate benzoxaboroles with broad spectrum activity
against diverse fungal plant pathogens. Through whole genome
sequencing of Saccharomyces cerevisiae isolates selected for stable
resistance to these compounds, we identified mutations in the protein prenylation-related genes, CDC43 and ERG20. Allele-
swapping experiments confirmed that point mutations in CDC43, which encodes an essential catalytic subunit within geranylgeranyl
transferase I (GGTase I) complex, were sufficient to confer resistance to the benzoxaboroles. Mutations in ERG20, which encodes an
upstream farnesyl pyrophosphate synthase in the geranylgeranylation pathway, also conferred resistance. Consistent with impairment
of protein prenylation, the compounds disrupted membrane localization of the classical geranylgeranylation substrate Cdc42. Guided
by molecular docking predictions, which favored Cdc43 as the most likely direct target, we overexpressed and purified functional
GGTase I complex to demonstrate direct binding of benzoxaboroles to it and concentration-dependent inhibition of its transferase
activity. Further development of the boron-containing scaffold described here offers a promising path to the development of GGTase
I inhibitors as a mechanistically distinct broad spectrum fungicide class with reduced potential for cross-resistance to antifungals in
current use.
subsistence farmers in developing countries, can be affected
by anthracnose caused by Colletotrichum species.5 In addition
to infectious diseases, contamination of foods by mycotoxins
can have serious consequences for human health. A prime
example of such contamination is aflatoxin, a carcinogenic
toxin produced by Aspergillus flavus. It is estimated that
consumption of aflatoxin-contaminated food may be respon-
sible for up to 155 000 cases of liver cancer per year.6 These
threats to food security and human health demand new
strategies to thwart plant pathogenic fungi.
INTRODUCTION
■
The fungal kingdom encompasses an estimated 2.2 to 3.8
million species, inhabiting nearly all environmental niches
around the globe.1 Of the diverse niches that fungi inhabit,
association with plants is particularly important, as fungi can
engage in symbiotic or parasitic relationships with agricultur-
ally important crops. Approximately 90% of land plants are
colonized by mycorrhizal fungi, which assist host plants by
improving phosphate and water uptake.2 In contrast, fungi are
also the causative agents of devastating diseases in staple crops
including wheat, rice, and maize, such that they pose a
significant threat to global food security. Each year, rice blast
caused by Magnaporthe oryzae leads to 10−30% harvest loss,
while emerging strains of Puccinia graminis can cause up to
100% loss in various wheat cultivars.2,3 In addition to cereal
grains, other staple crops of developing countries are also
susceptible to fungal pathogens. Bananas and plantains,
important staple crops of tropical countries, can be severely
affected by Mycosphaerella fijiensis, the causative agent of Black
Sigatoka.4 Sorghum and cassava, widely cultivated by
The primary approach to combating plant pathogenic fungi
has been widespread deployment of chemical fungicides in the
fields, which has been accompanied by increases in global crop
Received: April 15, 2020
Accepted: June 9, 2020
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a
Scheme 1. Synthesis of Benzoxaboroles
a
Reagents and conditions: (i) NaBH4, THF, 0 °C to rt; (ii) fuming HNO3, −30 °C; (iii) H2, Pd/C, EtOAc, rt; (iv) NCS, DMF, 0 °C to rt; (v)
thiophosgene, NaOH, DCM, 0 °C to rt; (vi) DMF, 80 °C; (vii) EtOAc, rt; (viii) (a) triphosgene, DIPEA, DCM, 0 °C to rt; (b) K2CO3, DCM, 40
°C.
yields over the past 40 years.7 There are four major classes of
modern fungicides: methyl benzimidazoles (MBCs) that target
β-tubulin, quinone inhibitors that target mitochondrial
cytochrome b, succinate dehydrogenase inhibitors, and
demethylation inhibitors, primarily azoles, that target 14α-
demethylase, an essential component of the ergosterol
biosynthetic pathway in fungi.7 Of these, azoles are one of
the most widely used fungicides in the fields due to their low
cost and broad-spectrum activity.8 It is estimated that
approximately half of the agricultural area for cereal and
grapevine are treated with azole fungicides in the European
Union.9 Inhibition of 14α-demethylase, encoded by CYP51
(also known as Erg11), by azoles causes accumulation of sterol
intermediates and depletion of ergosterol in the fungal cell
membrane, eventually leading to an arrest in fungal cell
growth.8 However, due to increased use in the fields, resistance
is observed to all of the fungicide classes described
above.8,10−12 For example, MBCs, one of the earliest single-
target fungicides introduced in the market to control cereal
eyespot disease caused by Oculimacula species, was discon-
tinued in many European countries in 1980s due to the
emergence of resistance.7,13 Azoles were introduced as a
replacement, which led to the emergence of azole resistance in
the field as early as the 1990s, with mutation in CYP51 as the
primary mode of resistance.8,13
Resistance to agricultural fungicides poses a severe threat to
human health beyond the issue of food security. The
fundamental problem arises from the widespread deployment
of the same class of antifungal agents in agriculture and
medicine, the primary example being azoles.14 It is clear that
mutations in CYP51 that emerge in fungi exposed to azoles in
the field confer cross resistance to azoles used to treat human
patients in the clinic.15,16 This is a pressing concern because
many human fungal pathogens are also common in the
environment, such as Aspergillus fumigatus, Cryptococcus
neoformans, and Histoplasma capsulatum, where they may
inhabit the soil or plants that come directly in contact with
agricultural fungicides.17−19 As such, the widespread use of
azole fungicides in agriculture can select for environmental
reservoirs of resistant fungal pathogens that may repeatedly
infect susceptible populations. In support of this concept, a
number of recent studies have reported that resistant clinical A.
fumigatus isolates recovered from azole naive patients likely
acquired their resistance as a result of exposure to azole
fungicides used in agriculture.20 Due to the increased incidence
of resistance to agricultural fungicides and the potential for
cross-resistance to clinically useful azole antifungal drugs, an
urgent need exists to develop new agricultural fungicides that
act upon cellular pathways distinct from those targeted by the
current armamentarium of clinical antifungals.
Post-translational modification of proteins with isoprenoid
lipids, collectively termed prenylation, mediates key protein−
protein and protein−membrane interactions in eukaryotic
cells, including fungi, suggesting prenylation could provide a
promising target for development of new agricultural
fungicides. Within the broader category of prenylation,
attachment of a 20-carbon isoprenoid lipid to the C-terminus
of proteins carrying a CaaL amino acid motif is specifically
termed geranylgeranylation. This modification is catalyzed by
geranylgeranyl transferase I (GGTase I), a heterodimeric
complex that is composed of Cdc43 and Ram2 in fungi.21 In
the model fungus Saccharomyces cerevisiae, geranylgeranylation
is essential, at least in part because it is required for proper
localization of essential GTPases involved in signal trans-
duction such as Cdc42.22 While GGTase I inhibitors have been
explored extensively as anticancer agents, investigation of
GGTase I as an antifungal target has only recently begun.23−25
In an effort to address the need for new classes of antifungals
with novel mechanisms of action, we synthesized a series of 6-
thiocarbamate benzoxaboroles with potent activity against a
broad panel of fungal plant pathogens. Boron-containing
compounds are proving a rich source of bioactive molecules
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Table 1. Antifungal Activity of Benzoxaboroles.
minimum inhibitory concentration (MIC95, μM)
compound ID
A. flavus
C. albicans
B. cinerea
M. fljiensis
R. solani
C. sublineolum
7
8
9
2.59 (0.017)
8.89 (0.020)
>40 (0.242)
0.71 (0.004)
2.53 (0.006)
>40 (0.196)
>40 (0.489)
>40 (0.329)
2.93 (0.008)
>40 (0.003)
20.72 (0)
1.29 (0.006)
8.89 (0.003)
>40 (0.140)
0.71 (0.008)
0.62 (0.003)
2.59 (0.033)
8.89 (0.006)
>40 (0.350)
2.93 (0.008)
10.17 (0.003)
2.59 (0.003)
17.78 (0.026)
>40 (0.007)
0.71 (0.004)
0.62 (0.003)
17.78 (0.003)
>40 (0.399)
5.87 (0.011)
5.07 (0.006)
chlorothalonil
tebuconazole
a
Sensitivity of fungal pathogens to growth inhibition by benzoxaboroles. Minimum concentration of compound (MIC) required to inhibit growth
by 95% was determined by averaging optical density (OD600) measurements from nine technical replicates. Standard deviation of the mean is
indicated in parentheses.
Figure 1. Mutations in either CDC43 or ERG20 within the protein geranylgeranylation pathway confer resistance to the benzoxaboroles. (A)
Sensitivity of an efflux pump-compromised strain of the model fungus S. cerevisiae to benzoxaboroles; the diploid parental strain harbors
homozygous deletion of SNQ2, PDR3, and PDR1. (left) Growth in liquid culture as measured by optical density (OD600) in the presence of a 2-fold
dilution series of the indicated compounds. Relative growth is presented in heat map format. Scale bar is provided below panel; bright green
represents full growth (1) and black represents no growth (0) relative to no-compound control wells. Each colored box represents the mean of
duplicate determinations. (right) After measuring relative growth, cells were spotted on compound-free YPD agar to determine their viability. The
entire experiment was repeated once with quantitatively similar results. (B) Sensitivity testing of benzoxaborole-resistant S. cerevisiae mutants
generated by in vitro selection. Relative growth is presented in heat map format as described for panel A. Mutations identified by whole genome
sequencing of the resistant isolates are indicated on the right. The experiment was repeated once with quantitatively similar results. (C) Sensitivity
testing of recombinant allele-swap mutants. ERG20 and CDC43 mutants and their respective hygromycin B-resistance marker-matched control
strains were generated in the same efflux pump-compromised S. cerevisiae parental strain used for selection experiments. Relative growth is
presented in heat map format as described for panel A. The experiment was repeated once with quantitatively similar results.
C
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with the potential to engage a wide spectrum of therapeutic
targets through both covalent and noncovalent interactions.26
FDA-approved boron-containing drugs include tavaborole, a
leucyl-tRNA synthethase inhibitor with antifungal activity, and
Eucrisa, a human phosphodiesterase 4 (PDE4) inhibitor.27,28
To identify potential target(s) responsible for the antifungal
activity of the new benzoxaborole series reported here, we used
the model fungus S. cerevisiae and sequenced the genomes of
benzoxaborole-resistant mutants. This analysis identified
Cdc43, the catalytic subunit of the geranylgeranyl transferase
I (GGTase I) complex in fungi, as the most likely candidate
target. We then leveraged cellular and biochemical assays to
confirm the compounds as new boron-containing GGTase I
inhibitors with activity against a broad spectrum of fungal plant
pathogens.
drug target genes and impair target engagement. We selected
for mutants resistant to 7 or 8 using the model yeast S.
cerevisiae due to its well-annotated genome and the diverse
genomic tools available for this organism.30 To enhance the
likelihood of identifying dominant gain-of-function mutations
directly relevant to compound mode of action rather than
noninformative mutations indirectly altering sensitivity
through effects on growth or compound efflux, a diploid,
drug-sensitized strain was used in which three ATP-binding
cassette (ABC) transporter genes were deleted: SNQ2, PDR3,
and PDR1.30 Both 7 and 8 inhibited growth of this sensitized
strain at 2.5 and 1.25 μM, respectively (Figure 1A). These
benzoxaboroles displayed fungicidal activity, as cells exposed to
concentrations above the MIC failed to grow when
subsequently spotted on drug-free medium (Figure 1A). Our
other benzoxaborole analog, 9, did not show any antifungal
activity.
RESULTS
■
Synthesis and Antifungal Spectrum of Benzoxabor-
oles. The preparation of analogues 7, 8, and 9 commenced
with synthesis of the key intermediate 6-amino-7-chloro
benzoxaborole 5 (Scheme 1). Reducing the formyl group of
2-formylphenylboronic acid 1 with sodium borohydride,
followed by spontaneous cyclization to construct the five-
membered oxaborole ring, produced benzoxaborole 2, which
was regioselectively nitrated at the 6-positon using fuming
nitric acid to afford 3. Hydrogenation of 3 provided 4, followed
by regioselective chlorination at the 7-position of 4 with N-
chlorosuccinimide (NCS) to give the key intermediate 5.
Treating 5 with thiophosgene in the presence of sodium
hydroxide afforded isothiocyanate 6, which condensed in situ
with (4-fluorophenyl) methanol or propane-2-thiol to afford
O-thiocarbamate 8 and dithiocarbamate 7, respectively.
Carbamate 9 was prepared by treatment of 2-methoxyethan-
1-ol with triphosgene in the presence of N,N-diisopropylethyl-
amine (DIPEA), followed by the key intermediate 5.29
The three benzoxaborole analogs, 7, 8, and 9, along with the
commonly used agricultural fungicides chlorothalonil and
tebuconazole, were tested for antifungal activity against a
panel of diverse fungi including plant pathogens, Botrytis
cinerea, Mycosphaerella fijiensis, Rhizoctonia solani, and
Colletotrichum sublineolum, as well as a common crop
contaminant, Aspergillus flavus, and the human fungal
pathogen Candida albicans (Table 1). The benzoxaboroles 7
and 8 were broadly active against plant pathogens but not C.
albicans. Minimum inhibitory concentrations (MIC95) were at
or below 20 μM, similar to conventional agricultural
fungicides. In contrast, compound 9 showed no activity up
to 40 μM.
We recovered seven independent resistant mutants, six of
which were from the selection with 7 and demonstrated 2−40-
fold increased resistance to this compound (Figure 1B); One
mutant was recovered from selection with 8 and had ≥32-fold
increased resistance. All mutants demonstrated cross-resistance
to both bioactive benzoxaboroles (Figure 1B), suggesting that
the compounds likely shared a common mode of action.
To identify candidate resistance mutations, we performed
whole genome sequencing of resistant isolates. Comparing the
genome sequence of our seven resistant isolates to that of the
parental strain, we identified heterozygous mutations CDC43/
CDC43C1083G (four isolates), CDC43/CDC43T509G (one
isolate), and ERG20/ERG20A284C (two isolates), which caused
following substitutions: Cdc43C361W, Cdc43I170S, and Erg20Y95S
,
respectively (Figure 1B). To determine whether these
alterations alone were sufficient to confer resistance, we
expressed each mutant allele in the parental diploid strain and
confirmed that each acted in a dominant fashion to confer
resistance (Figure 1C). Mutant ERG20 conferred similarly
robust resistance to both 7 and 8, while CDC43 mutations
conferred greater resistance to 7 than 8 when comparing
strains expressing mutant alleles to their respective hygromycin
B-resistance marker-matched control strain with wild-type
alleles.
As a complementary approach to assessing the relevance of
CDC43 and ERG20 to the mode of action of our
benzoxaboroles, we determined whether reduced dosage of
either gene might confer hypersensitivity to the compounds, as
would be expected for a gene encoding a compound’s target or
components of pathways involving the target. We found that
strains carrying heterozygous deletions of either gene
(CDC43/Δ or ERG20/Δ) were hypersensitive to 7 and 8
compared to the parental strain, further implicating Cdc43 and
Erg20 in the mode of action of these benzoxaboroles
(Supplementary Figure 2).
In S. cerevisiae, both Cdc43 and Erg20 are required for a
range of essential metabolic processes. Cdc43 heterodimerizes
with the adapter protein Ram2 (RAS protein and α-factor
maturation) to form GGTase I complex (GGTase I),21,22
whereas Erg20 catalyzes the production of farnesyl-pyrophos-
phate (F-PP), a 15-carbon intermediate required for diverse
biosynthetic processes including protein farnesylation, produc-
tion of sterols, and generation of the geranylgeranyl-
pyrophosphate (GG-PP), which is utilized downstream by
Cdc43.31
The three benzoxaboroles were also tested for their chemical
stability in phosphate buffer (pH 7.4) at RT over 72 h
(Supplementary Figure 1A). Compounds 8 and 9 remained
intact over the entire incubation period. Compound 7 was
much less stable, with a reduction of 75% in parent compound
within 24 h and near complete loss by 72 h. When we tested
compound stability under the more acidic conditions (pH 5.0)
characteristic of the culture medium used to perform MIC
assays, the stability of compound 7 was markedly improved. As
expected, the stability of compound 9 showed no change
(Supplementary Figure 1B).
Mutations in CDC43 and ERG20 Confer Resistance to
Benzoxaboroles. Selection for drug-resistant mutants
provides a powerful approach to elucidate the mode of action
of bioactive compounds as resistance mutations often arise in
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To assess the potential for direct interaction of our
benzoxaboroles with either of the proteins encoded by our
genetic hits, we first generated a homology model for the
structure of S. cerevisiae GGTase I using a published C. albicans
structure32 as the template. Docking experiments with 7 and 8
predicted the compound to occupy the active site with its
boron group interacting with the catalytic zinc(II) ion and
Lys331 in the negatively charged tetrahedral form, while its
hydrophobic 6-thiocarbamate substituent fills the hydrophobic
substrate site (Figure 2). Residues C361 and I170 have not
residues in prokaryotic homologues are important in
determining the chain length of the prenyl product produced
by this enzyme. Substitution of native tyrosine with either
serine or histidine has been shown to favor the production of
GG-PP instead of F-PP by Erg20. This insight suggests that the
Y95S mutation might help overcome downstream inhibition of
Cdc43 by 7 and rescue the compound’s toxicity by increasing
GG-P abundance and driving GGTase I activity.31,34,35
Consistent with Cdc43 as the most likely direct target of 7,
this compound’s lack of activity against C. albicans (Table 1)
could be explained by the fact that Cdc43 is not an essential
gene in this fungus.36 Overall, our genetic data and structural
modeling support inhibition of protein geranylgeranylation as
the primary mode of action for our antifungal benzoxaboroles.
Importantly, GGTase I is not essential for viability or normal
development in the model plant, Arabidopsis thaliana,
conferring an important advantage to targeting geranylger-
anylation in the development of crop fungicides.37
Benzoxaboroles 7 and 8 Disrupt Membrane Local-
ization of Cdc42 in S. cerevisiae. Geranylgeranylation is
required for proper localization of essential signaling proteins,
such as the Rho-family GTPase Cdc42. Disruption of GGTase
I function or mutation of the CaaL recognition site on the C-
terminus of Cdc42 impairs its proper membrane local-
ization.22,38 To determine if our antifungal benzoxaboroles
impair normal membrane localization of Cdc42, we con-
structed a centromeric vector to express GFP or a GFP−
Cdc42 fusion protein under control of the MET17 promoter,
transformed into S. cerevisiae strain BY4742, and monitored
their intracellular localization by fluorescence microscopy. As
expected, GFP showed diffuse cytoplasmic localization while
GFP−Cdc42 showed the expected intracellular membranous
localization (Figure 3). Addition of a cell cycle arresting agent,
hydroxyurea, did not alter the localization pattern of GFP−
Cdc42. However, when the strain expressing GFP−Cdc42 was
treated with 7 or 8, the fluorescence localization pattern shifted
from membranous to cytoplasmic. Taken together, these
results suggest that benzoxaborole treatment impairs GFP−
Cdc42 localization, consistent with inhibition of GGTase I
activity by the compounds.
Benzoxaboroles 7 and 8 bind to GGTase I Complex
and Inhibit Its Activity In Vitro. Next, we turned to
biochemical approaches to determine if the benzoxaboroles 7
and 8 directly inhibit GGTase I activity. We C-terminally
tagged Cdc43 with 6XHis via a Tobacco Etch Virus (TEV)-
linker sequence and placed both this tagged protein and Ram2
under the control of the GAL1 galactose-inducible promoter
using independent 2 μm plasmids with distinct markers that
enable selection for coexpression in S. cerevisiae. We trans-
formed the two plasmids in S. cerevisiae strain, BY4741, then
verified the functionality of Cdc43-6XHis and Ram2 by
showing that coexpression of these tagged proteins was
sufficient to confer resistance to the benzoxaboroles (Supple-
mentary Figure 3A).
Figure 2. In silico homology modeling and docking analysis predicts
novel benzoxaborole interactions with GGTase I: (top) compound 7
and (bottom) compound 8 (color code: boron, pink; carbon,
magenta, oxygen, red; nitrogen, deep blue; sulfur, yellow; chlorine,
green; fluorine, light blue) occupy the active site of S. cerevisiae
GGTase I (color code: carbon, cyan; oxygen, red; nitrogen, blue;
sulfur, yellow) directly interacting with C361 and I170.
been previously reported as important in the catalytic function
of GGTase I, but our docking model highlights a role for them
in enhancing inhibitor binding by interacting directly with the
isopropyl group of 7 or fluorobenzyl group of 8 away from the
catalytic site. The substitutions we observed in selection
experiments, C361W and I170S, would be expected to weaken
the binding of the inhibitor by narrowing the binding pocket or
reducing the hydrophobic packing interactions for both the
isopropyl group and fluorobenzyl group of the inhibitors
(Figure 2).
We also generated a homology model of S. cerevisiae Erg20
based on a previously reported human crystal structure and
attempted to dock 7 into this model.33 Although two potential
sites for binding of 7 were identified, docking at either did not
explain how a Y95S substitution could cause resistance to 7.
Interestingly, Y95 of Erg20 in humans or the corresponding
To perform biochemical assays, the GGTase I complex was
purified from whole yeast extracts using nickel resin, followed
by TEV protease treatment to remove the 6XHis tag. Purity of
the final GGTase I preparation was determined to be
approximately 50% based on SDS-PAGE (Supplementary
Figure 3B). Using this affinity-purified material, we first
examined the potential binding of 7 and 8 to GGTase I
complex using a well-established thermal denaturation
approach, which is based on the ability of a ligand to stabilize
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Figure 3. Benzoxaborole treatment causes mislocalization of GFP−Cdc42 fusion protein within cells. Cells were grown overnight in SC-HIS then
subcultured in SC-HIS-MET for 6 h in the absence or presence of compounds. Cells were imaged with DIC and GFP channels on Apotome. Scale
bar represents 10 μm.
the conformation of the protein to which it is bound and shift
the temperature at which it melts.39 We incubated 5 μM
GGTase I complex with test compounds (200 μM) and
measured changes in fluorescence intensity over a temperature
ramp from 25 to 98.6 °C. Consistent with their binding to the
GGTase I complex, 7 and 8 markedly shifted the melting
curves of GGTase I complex by 5.8 and 6.4 °C, respectively,
compared to solvent (DMSO) control (Figure 4A).
Furthermore, our inactive benzoxaborole, 9, had a more
modest effect on the melting temperature of the GGTase
complex (2.4 °C). We also tested whether the mammalian
GGTase I inhibitor, GGTI-2133, binds to fungal GGTase I by
thermal denaturation assay.40 GGTI-2133 is a class of
peptidomimetic inhibitor, which engages the substrate
recognition site of GGTase I by mimicking the CaaL
recognition sequence.41 Although we did not observe any
growth inhibition for this compound in S. cerevisiae, GGTI-
2133 shifted the melting temperature of the purified complex
(4 °C), suggesting that it does interact with fungal GGTase I in
vitro as would be expected given conservation of the CaaL
recognition motif between fungi and humans (Figure 4A,
Supplementary Figure 4).
Having obtained evidence for binding, we proceeded to
determine whether the benzoxaboroles could directly inhibit
GGTase I function, as measured by covalent labeling of a
canonical target protein (RhoA) with NBD-FPP, a fluorescent
lipid donor that mimics the geranylgeranyl moiety.42 Following
labeling reactions in the presence or absence of test
compounds, relative transferase activity was measured by
resolving reaction mixtures using SDS-PAGE and quantitating
fluorescently labeled RhoA by imaging the gels. Consistent
with our thermal shift data, 7, 8, 9, and GGTI-2133 all caused
concentration-dependent inhibition of RhoA labeling, indicat-
ing that all four compounds inhibit fungal GGTase I activity
(Figure 4B). Plotting relative fluorescence intensity as a
function of compound concentration, we found that 7 and 8
were more potent than 9 and GGTI-2133 (Figure 4C).
All mechanistic studies were performed in the model yeast S.
cerevisiae. Although the GGTase I complex is highly conserved
among fungi, we acknowledge the formal possibility that the
activity of 7 and 8 demonstrated against a panel of
agriculturally important fungi (Table 1) could be mediated
through a mechanism other than inhibition of the GGTase I
complex. In addition, we found that growth of the human
fungal pathogen C. albicans is not inhibited by compounds 7
and 8; most likely because GGTase I function is not essential
in this organism.36 However, in the era of escalating
antimicrobial resistance in patients, targeting nonessential
processes that affect a pathogen’s drug susceptibility or
virulence have been gaining substantial attention as it could
reduce the selection for drug resistance.43,44 In C. albicans, a
GGTase I inhibitor, L-269289, was reported to potentiate the
activity of the echinocandin antifungal caspofungin. In this
organism, CDC43 or RAM2 expression was also found essential
for virulence in a mouse model of disseminated infection.24,25
These findings suggest that despite their lack of fungicidal
activity against this pathogen, boron-containing GGTase I
inhibitors might still provide a useful approach to the
management systemic C. albicans infections in humans.
All three compounds reported here share the identical
benzoxaborole group linked to structurally related carbamate,
thiocarbamate, or dithiocarbamate groups (Scheme 1). Our in
silico modeling predicts that the dithio- or thiocarbamate group
both functions as a structural linker connecting two essential
pharmacophores and also interacts with the hydrophobic
pocket of GGTase I itself. Although no benzoxaborole
GGTase I inhibitors have been previously reported, this class
of compounds has proven a rich source of useful human
therapeutics from antifungal drugs to anti-inflammatory
agents.27,28 In support of drug-development efforts, struc-
ture−activity relationship (SAR) studies with 6-substituted
F
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Figure 4. Benzoxaboroles bind fungal GGTase I complex and inhibit its activity. (A) Thermal shift assay to assess binding of purified GGTase I
complex isolated from S. cerevisiae to benzoxaboroles and the mammalian GGTase I inhibitor GGTI-2133. The negative first derivative of dye
fluorescence/temperature is plotted versus temperature over a thermal ramp from 25 to 100 °C for each of the indicated compounds (100 μM).
The colored numbers indicate changes in the first derivative of fluorescence between DMSO control and the corresponding compound treatment
conditions. (B) Transfer of fluorescent NBD-FPP label to the model geranylgeranylation substrate RhoA. Purified GGTase I complex was
incubated with human recombinant RhoA and NBD-FPP as a geranylgeranyl pyrophosphate mimetic. Reactions were performed in the absence or
presence of inhibitors at 30 °C for 2 h. Reactions were quenched by boiling in loading buffer, and samples were fractionated under denaturing
conditions by SDS-PAGE. Green fluorescence of the NBD-FPP label was visualized by scanning imager and then total RhoA was visualized by
Coomassie blue staining of the gel. Pseudocolors were added for clarification. (C) Quantification of fluorescence. EGFP and Coomassie blue
intensities were quantified by ImageJ. EGFP intensities were calculated relative to the Coomassie blue intensities, and then relative EGFP
intensities were calculated to the DMSO sample.
benzoxaboroles have been performed across a wide variety of
targets and made clear the importance of modifications to the
6-substitution for improving bioactivity either by increasing
target affinity or by altering lipophilicity to decrease
susceptibility to drug efflux pumps.45−47 Here, we observed
major differences in bioactivity, chemical stability, and target
engagement and inhibition based on variation in substitutions
at the 6-position for our benzoxaborole compounds.
Carbamate containing compound 9 showed no antifungal
activity, while dithiocarbamate containing compound 7
showed reduced chemical stability in aqueous buffer at pH
7.4. Carbamate containing compound 9 did not demonstrate
antifungal activity in culture at concentrations up to 40 μM,
even though the compound is stable and target engagement
assays indicate that the compound can inhibit GGTase I
function to some degree. Whether the lack of antifungal
activity for 9 is solely due to reduced target affinity or if
additional factors such as poor fungal permeability or
susceptibility to efflux pumping play a role remains unclear.
Although dithiocarbamate containing compound 7 displayed
poor stability in aqueous solution at pH 7.4, its fungicidal
activity and in vitro target engagement to GGTase I complex at
low micromolar concentrations suggest that it could still be
useful for agricultural purposes. In fact, decomposition over
time could be an asset in the field to avoid undesirable
persistence and downstream environmental contamination.
Furthermore, thiocarbamate containing compound 8 provides
an alternate option when stability is desired without loss in
antifungal activities. Overall, the 6-thiocarbamate benzoxabor-
ole series described here should provide a tractable, well-
supported route to the design and synthesis of diverse analogs
for future SAR studies focusing on inhibition of fungal GGTase
G
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= 10/1) to give 5 (132 mg, 0.72 mmol, 11% yield) as a yellow solid.
1H NMR (DMSO-d6, 400 MHz): δ 8.89 (s, 1 H), 7.03 (d, J = 8.0 Hz,
1 H), 6.92 (d, J = 8.0 Hz, 1 H), 5.21 (s, 2 H), 4.82 (s, 2 H) ppm. MS
(ESI): mass calcd for C7H7BClNO2 183.03, m/z found 184.2 [M +
H]+. Purity by HPLC: 96.33% (220 nm), 91.48% (254 nm).
I and using both biochemical and whole cell end points.
Indeed, further exploration of the 6-thiocarbamate benzox-
aborole scaffold appears to offer a promising path to the
development of much needed broad-spectrum agricultural
fungicides with reduced potential for driving cross-resistance to
antifungals currently in use for medical and veterinary
purposes.
7-Chloro-6-isothiocyanatobenzo[c][1,2]oxaborol-1(3H)-ol
(6). To a solution of 6-amino-7-chlorobenzo[c][1,2]oxaborol-1(3H)-
ol (1 g, 5.4 mmol) and thiophosgene (1.23 mL, 16.2 mmol) in
anhydrous DCM (8 mL) was added sodium hydroxide (0.65 g, 16.2
mmol) in portions at 0 °C under N2 atmosphere; then the mixture
was warmed to RT. The reaction mixture was cooled back to 0 °C, its
pH was adjusted to 5 by adding 1 N HCl, and the resulting mixture
was extracted with EtOAc (2 × 25 mL). Combined organic phase was
washed with H2O and brine, dried over anhydrous Na2SO4, and
filtered. The filtrate was concentrated in vacuo to afford the
EXPERIMENTAL SECTION
■
General Methods. Commercial reagents and solvents were used
as supplied. Low resolution mass spectroscopy was carried out using
liquid chromatography−mass spectrometry (LC/MS) on an Agilent
1
instrument using electrospray ionization (ESI). H NMR magnetic
resonance spectra were obtained on a Bruker instrument in CDCl3 or
DMSO-d6 at 400 MHz and 298 K unless otherwise noted. Chemical
shifts were expressed in ppm relative to an internal standard,
tetramethylsilane (ppm = 0.00). The following abbreviations were
utilized to describe peak patterns when appropriate: br = broad, s =
singlet, d = doublet, and m = multiplet. Compounds used in biological
studies had purities that were >95%, determined by HPLC based on
ultraviolet detection at 210 and 254 nm and 1H NMR
(Supplementary Figure 5).
1
intermediate 6 (1.1 g, yield 91%) as a pale yellow solid. H NMR
(300 MHz, DMSO-d6): δ 9.38 (s, 1H), 7.61 (d, J = 8.0 Hz, 1H), 7.42
(d, J = 8.0 Hz, 1H), 4.99 (s, 2H). MS (ESI): mass calcd for
C8H5BClNO2S 224.98, m/z found 224.0 [M − H]−. Purity by HPLC:
97.25% (220 nm), 99.08% (254 nm).
O-(4-Fluorobenzyl)(7-chloro-1-hydroxy-1,3-dihydrobenzo-
[c][1,2]oxaborol-6-yl)carbamothioate (8). A solution of 7-chloro-
6-isothiocyanatobenzo[c][1,2]oxaborol-1(3H)-ol (300 mg, 1.3
mmol) and (4-fluorophenyl)methanol (0.3 mL, 2.8 mmol) in DMF
(0.5 mL) was stirred at 80 °C for 4 h. The solvent was removed in
vacuo to give a residue, which was purified by Prep-HPLC to give 8
Benzoxaborole (2). To a solution of (2-formylphenyl)boronic
acid (10 g, 66.69 mmol, 1 equiv) in THF (100 mL) was added
NaBH4 in portions (5.05 g, 133.39 mmol, 2 equiv) at 0 °C over 5
min. After addition, the reaction mixture was kept stirring at 25 °C for
1 h. The reaction was then quenched by adding aq. HCl (2 M) at 0
°C until pH = 7. The resulting mixture was diluted with EtOAc (30
mL) and extracted with EtOAc (30 mL × 3). The combined organic
layers were washed with brine (30 mL × 3), dried over anhydrous
Na2SO4, filtered, and concentrated under reduced pressure to give 2
1
(59 mg, 13%) as a white solid. H NMR (400 MHz, DMSO-d6): δ
10.94 (br s, 1H), 9.19 (s, 1H), 7.61−7.08 (br. m, 6H), 5.49 (s, 2H),
4.99 (s, 2H) ppm (Supplementary Figure 5). HPLC purity: 98.7% at
210 nm and 99.5% at 254 nm. MS (ESI): mass calcd for
C15H12BClFNO3S 351.03, m/z found 350.0 [M − H]−.
Isopropyl(7-chloro-1-hydroxy-1,3-dihydrobenzo[c][1,2]-
oxaborol-6-yl)carbamodithioate (7). The title compound was
prepared from propane-2-thiol and 7-chloro-6-isothiocyanatobenzo-
[c][1,2]oxaborol-1(3H)-ol using the procedure employed for the
synthesis of O-(4-fluorobenzyl)(7-chloro-1-hydroxy-1,3-
dihydrobenzo[c][1,2]oxaborol-6-yl)carbamothioate. It was obtained
1
(7.7 g, 57.49 mmol, 86.2% yield) as a white solid. H NMR (CDCl3,
400 MHz): δ 7.78 (d, J = 7.2 Hz, 1 H), 7.53−7.49 (m, 1 H), 7.41−
7.36 (m, 2 H), 5.75 (s, 1 H) and 5.15 (s, 2 H) ppm.
1-Hydroxy-6-nitro-3H-2,1-benzoxaborole (3). To a solution of
fuming HNO3 (70 mL) was added benzo[c][1,2]oxaborol-1(3H)-ol
(6.7 g, 50.02 mmol, 1 equiv) at −30 °C over 5 min; the resulting
mixture was stirred at −30 °C for 30 min. Then the mixture was
poured into water and ice mixture (80 mL) and kept stirring at 0 °C
for 40 min. The precipitate formed from the reaction mixture was
collected by filtration and dried in vacuo to afford 3 (6 g, 33.53 mmol,
67.0% yield, 100% purity) as a pale yellow solid. 1H NMR (DMSO-d6,
400 MHz): δ 9.59 (s, 1H), 8.57 (s, 1H), 8.33 (dd, J = 8.4, 2.0 Hz, 1
H), 7.69 (d, J = 8.4 Hz, 1 H), 5.12 (s, 2 H) ppm. MS (ESI): mass
calcd for C7H6BNO4 178.94, m/z found 180.1 [M + H]+. Purity by
HPLC: 100% (220 nm) and 100% (254 nm).
1
as a white solid (yield 22%). H NMR (400 MHz, DMSO-d6): δ
11.39 (s, 1H), 9.24 (s, 1H), 7.47 (d, J = 7.9 Hz, 1H), 7.40 (d, J = 7.9
Hz, 1H), 5.02 (s, 2H), 3.94−3.87 (m, 1H), 1.35 (d, J = 5.7 Hz, 6H)
ppm (Supplementary Figure 5). HPLC purity: 97.9% at 210 nm and
99.1% at 254 nm. MS (ESI): mass calcd for C11H13BClNO2S2 301.02,
m/z found 301.9 [M + H]+.
2-Methoxyethyl(7-chloro-1-hydroxy-1,3-dihydrobenzo[c]-
[1,2]oxaborol-6-yl)carbamate (9). To a solution of 2-methox-
yethan-1-ol (760 mg, 10.0 mmol) and N,N-diisopropylethylamine
(516 mg, 4.0 mmol) in DCM (25 mL) was added a solution of
triphosgene (1.2 g, 4.0 mmol) in DCM (10 mL) dropwise at 0 °C.
The resulting mixture was stirred at 0 °C for 30 min, then warmed to
RT and kept stirring overnight. 6-Amino-7-chlorobenzo[c][1,2]-
oxaborol-1(3H)-ol (250 mg, 1.40 mmol) and K2CO3 (4.14 g, 30
mmol) were added into the reaction mixture, and the resulting
reaction mixture was heated to 40 °C and kept stirring for 30 min.
The reaction mixture was cooled to 0 °C, quenched by addition of 5
mL of H2O, neutralized by 6 N HCl to pH ≈ 3, and extracted with
EtOAc (2 × 50 mL). The combined organic phase was washed with
H2O and brine, dried over anhydrous Na2SO4, and filtered. The
filtrate was concentrated in vacuo to give a residue, which was purified
by prep-HPLC to give 9 (19 mg, yield 5%) as a white solid. 1H NMR
(400 MHz, DMSO-d6): δ 9.13 (s, 1H), 9.09 (s, 1H), 7.60 (d, J = 8.1
Hz, 1H), 7.33 (d, J = 8.0 Hz, 1H), 4.96 (s, 2H), 4.21−4.15 (m, 2H),
3.58−3.54 (m, 2H), 3.28 (s, 3H) ppm (Supplementary Figure 5).
HPLC purity: 98.35% at 210 nm and 96.95% at 254 nm. MS (ESI):
mass calcd for C11H13BClNO5 285.06, m/z found 286.3 (M + H)+.
Culture Conditions. Unless otherwise specified, plant pathogens
were maintained on potato dextrose agar (PDA) or V8 agar, and
spores were isolated from the cultures after 1−2 weeks of incubation
at RT (20−22 °C) with a 12-h fluorescent light (Philips, F40LW) and
12-h blacklight (Philips, F40T12) photoperiod. The final concen-
trations of all inocula were 5 × 104 CFU/mL. All yeast strains were
6-Aminobenzo[c][1,2]oxaborol-1(3H)-ol (4). To a solution of
1-hydroxy-6-nitro-3H-2,1-benzoxaborole (950 mg, 5.31 mmol, 1
equiv) in EtOAc (20 mL) was added Pd/C (200 mg, 10%) under
N2 atmosphere. The suspension was degassed under reduced pressure
and purged with H2 several times. The resulting mixture was stirred
under H2 atmosphere (15 psi) at 25 °C for 2 h. The reaction mixture
was filtered, and the filtrate was concentrated in vacuo to give a crude
product, which was purified by recrystallization from 20 mL of MTBE
to give 4 (0.73 g, 4.90 mmol, 92.3% yield, 100% purity) as a pale
1
yellow solid. H NMR (DMSO-d6, 400 MHz): δ 8.90 (s, 1 H), 7.03
(d, J = 8.0 Hz, 1 H), 6.87 (s, 1 H), 6.69 (d, J = 8.0 Hz, 1 H), 4.97 (s, 2
H), 4.80 (s, 2 H) ppm. MS (ESI): mass calcd for C7H8BNO2 149.06,
m/z found 150.1 [M + H]+. Purity by HPLC: 100% (220 nm) and
100% (254 nm).
6-Amino-7-chlorobenzo[c][1,2]oxaborol-1(3H)-ol (5). To a
solution of 6-aminobenzo[c][1,2]oxaborol-1(3H)-ol (1 g, 6.71 mmol,
1 equiv) in DMF (15 mL) was added N-chlorosuccinimide (NCS;
0.95 g, 7.11 mmol, 1.06 equiv) at 0 °C. The mixture was stirred at 25
°C for 1.5 h. The reaction was quenched by adding ice−water (50
mL) at 0 °C and extracted with EtOAc (10 mL × 3). The combined
organic layers were washed with brine (30 mL × 3), dried over
anhydrous Na2SO4, filtered, and concentrated in vacuo to give a
residue, which was purified by prep-TLC (SiO2, ethyl acetate/MeOH
H
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archived in 25% glycerol and stored at −80 °C. Yeast strains were
grown in YPD (1% yeast extract, 2% peptone, 2% glucose) at 30 °C
unless otherwise specified. For propagation of plasmids with
hygromycin B marker, yeast strains were grown in YPD plus 600
μg/mL hygromycin B. For propagation of plasmids with auxotrophic
marker, yeast strains were grown in synthetic complete (SC) medium
(0.674% yeast nitrogen base, 2% glucose) lacking specific amino acids
or nucleobase.
Selection of Resistant Mutants. Cell densities of overnight
cultures were quantified by hemocytometer counts. Approximately 1
× 108 cells were plated on YPD agar containing 40 μM compound 7
or 8 and incubated at 30 °C for 48 h.
Yeast Strain Construction. Strains used in this study and details
regarding their construction are provided in Supporting Information.
To prepare yeast cultures for transformation, cells were grown as 10
mL cultures until the optical density (OD600) was between 0.3 and
0.6. Cells were pelleted and resuspended in 60 μL of 100 mM lithium
acetate, followed by 15 min incubation at 30 °C. Cells were mixed
with 10 μL of linear DNA or 1 μL of plasmid DNA with 10 μL of
boiled salmon sperm DNA (10 mg mL−1) and 300 μL of PEG
mixture (40% poly(ethylene glycol), 100 mM lithium acetate) and
incubated at 30 °C for 30 min followed by 42 °C for 20 min. For
antibiotic markers, cells were recovered in 10 mL of YPD at 30 °C for
4 h before plating on YPD agar plus 600 μg/mL hygromycin B. For
auxotrophic markers, cells were directly plated on appropriate
synthetic SC medium lacking specific amino acids or nucleobase.
Plates were incubated at 30 °C for 48 h.
Plasmid Construction. Cloning procedures were performed
following standard protocols. Transformed DH5α competent
Escherichia coli cells (Invitrogen) were grown on LB with 2% agar
(Sigma) containing 100 μg/mL ampicillin and incubated at 37 °C
overnight. Plasmids and oligonucleotides used in this study are listed
in Supporting Information.
Antifungal Sensitivity Testing. S. cerevisiae. Approximately
1000 cells/well were inoculated with a 2-fold gradient of compounds
in 384-well microtiter plates to reach the final volume of 40 μL in
appropriate medium. The plates were incubated at 30 °C for 72 h
before measurement of the OD600 was performed using SpectraMax
M2e (Molecular Devices). Relative growth values were calculated by
normalizing OD600 measurements against the no-compound control
and were plotted as a heat map using R. To determine cell viability,
after reading OD600, cultures were spotted to YPD agar and incubated
at 30 °C for 24 h before imaging.
and 0.20 μg/mL]; control studies with 0 μg/mL of compounds were
performed in parallel for each plate). Plates sealed with clear polyester
film (VWR) were incubated at a temperature of ∼22 °C. The progress
of fungal growth was monitored at 72 h; yeast growth was monitored
at 48 h. The MICs were determined as the lowest antifungal
concentrations that inhibited fungal growth by greater than 95%
(determined as relative absorbance using the Bio-Tek Synergy H1
microplate reader at 600 nm) relative to the corresponding antifungal-
free control.
Compound Chemical Stability Testing. The buffers used in
this study consisted of 100 mM, pH 5, acetic acid/sodium acetate
buffer and 20 mM, pH 7.4, phosphate buffer. A 10 mM stock in
DMSO was prepared for all three compounds. To a sample vial, 20 μL
of compound stock was added to 380 μL of buffer for a final
incubation concentration of 500 μM. Each sample vial was vortexed
for approximately 30 s before collecting the initial data point. The
samples were incubated at RT, and data points were collected at T =
0, 1, 24, 48, and 72 h for analysis by HPLC on a Zorbax SB-C18
column (2.1 mm × 50 mm, 1.8 μM). Relative parent compound
concentration in each sample was determined by area under the peak
based on UV detection at 254 nm. Replicate samples for each
condition were analyzed (2 for incubations at pH 5 and 3 for
incubations at pH 7.4).
Whole Genome Sequencing. Overnight cultures of resistant
mutants were used to isolate genomic DNA for whole genome
sequencing. DNA was prepared using the PureLink Genomic DNA
Mini Kit (Invitrogen) according to the manufacturer’s instructions.
Sequencing libraries were prepared using the Nextera XT Kit
(Illumina) according to the manufacturer’s instructions. Libraries
were sequenced on the Illumina NextSeq platform using paired reads
(300 bp). The sequence reads were de-multiplexed and trimmed to
remove bases with Phred scores < Q30. Reads were aligned to S288C
reference genome using Bowtie2,48 and the alignment was visualized
using Integrative Genomics Viewer.49 MuTect was used to identify
unique mutations in resistant mutants.50
Haploinsufficiency Testing. Approximately 1000 cells/well of
appropriate heterozygous deletion strains in YPD medium were
incubated with a single compound concentration in 96-well format
(final volume 200 μL/well) and sealed with Breathe-Easy Sealing Film
(Diversified Biotech). The plates were incubated at 30 °C in an
Infinite 200 PRO plate reader (TECAN), and OD600 was measured
every 15 min for 48 h. Growth curve based on OD600 was plotted
using R.
Mycosphaerella fijiensis. Mycelial cultures of M. fijiensis isolates
11CR-33 grown on PDA medium were macerated in water, and 1−5
mL of the resulting suspension was pipetted onto plates of V8
medium. Cultures were incubated at 20 °C under continuous, cool-
white fluorescent and black light. After 5−7 days, sporulation plates
were stimulated to produce conidia by adding 2 mL of water, brushing
the plates with a paint brush or cell spreader, and removing the
resulting suspension. After another 5−7 days, conidia were harvested
in the same way, adding 2 mL of 0.01% Tween 20 solution, brushing
the plates to dislodge spores, and removing the spore suspension by
pipetting. Spores were diluted in half strength broth medium.
Rhizoctonia solani. Due to insufficient spore number obtained
from these fungi, inocula were prepared as mycelium fragments. In
brief, fungal mycelia grown on agar media were cut into 1 × 1 mm2
pieces and cultured in autoclaved broth medium (such as PDB and
V8). After 3−7 days of incubation at 22−24 °C, mycelia were
harvested by filtering through one layer of Miracloth. The mycelia
were homogenized in half strength of broth medium using a
household blender for 10 s and filtered through one layer Miracloth.
The resultant visible fragments were diluted in half strength broth
medium.
Molecular Docking. Sequence of S. cerevisiae Cdc43 was obtained
from UniProtKB (accession ID P18898). The crystal structure of C.
albicans protein geranylgeranyl transferase-I (PDB code 3DRA) was
used as a template for homology modeling. Sequences were aligned
and homology models were built using maestro Prime51 (Schrodinger,
LLC). Benzoxaborole inhibitors were then docked into the prepared
active site using Glide with fixed protein. The final docking model was
obtained by minimizing the complex using a water solvation model
and Optimized Potentials for Liquid Simulations 2005 force field
while maintaining a fixed protein backbone.
GFP−Cdc42 Fluorescence Microscopy. Cells were grown
overnight in 5 mL of SC-HIS at 30 °C. Overnight cultures were
washed and diluted to OD600 of 0.1 in 5 mL of SC-HIS-MET in the
absence or presence of 400 mM hydroxyurea or 2.5 μM
benzoxaboroles and were incubated at 30 °C for 6 h. Cells were
imaged on DIC and GFP channels on Inverted Axiovert 200M
Apotome imaging system (Zeiss).
Purification of the Fungal GGTase I Complex. Cells were
grown overnight in SC-HIS-LEU with 2% raffinose at 30 °C.
Overnight cultures were diluted to OD600 of 0.3 in 1 L of SC-HIS-
LEU with 2% galactose and 300 μM ZnSO4 to induce expression of
Cdc43-6XHis and Ram2 and incubated at 30 °C for 8 h. Cells were
pelleted and lysed using a French press in 20 mL of buffer A (25 mM
HEPES, pH 7.5, 40 mM NaCl, 20 μM ZnCl2, 2 mM DTT, 2 mM
MgCl2) with 1 mM PMSF and cOmplete Protease Inhibitor Cocktail
Tablet (Roche). Crude lysate (CL) was centrifuged at 20 000g at 4 °C
for 10 min. The pellet (P) was discarded. Clarified supernatant (S)
The MICs for individual compounds were determined by following
a modified broth microdilution protocol. The studies were performed
in flat bottom, 96-well microtiter plates (Greiner Bio-One). The
individual MICs were determined in triplicate in a final volume of 0.2
mL/well with antifungal concentrations of 0.2−25 μg/mL (8 serial
dilutions down from 25 μg/mL [25, 12.5, 6.25, 3.13, 1.56, 0.78, 0.39,
I
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was applied to 1 mL of Ni-NTA Agarose (Invitrogen) and incubated
at 4 °C for 30 min. The flow-through was discarded (FT). The resin
was washed with 10 mL of buffer A plus 10 mM imidazole (W10) and
then 10 mL of buffer A plus 25 mM imidazole (W25). The GGTase I
complex was eluted with 10 mL of buffer A plus 250 mM imidazole
(W250). Imidazole concentration was reduced back to 10 mM by
concentrating the purified protein with a 30 kDa MWCO Poly(ether
sulfone) filter (GE Healthcare) and washing with buffer A plus 10
mM imidazole two times (W250c). The GGTase I complex was
incubated with 50 units of AcTEV Protease (Invitrogen) at 4 °C
overnight. GGTase I complex was reapplied to 1 mL of Ni-NTA
Agarose, and the flow-through was collected (Post-TEV FT). The
flow-through was concentrated with Vivaspin 20, 30 kDa MWCO
Poly(ether sulfone) filter (GE Healthcare), and washed 2 times with
buffer A (Post-TEV FTc). The purified GGTase I complex was
aliquoted, flash frozen with liquid nitrogen, and stored at −80 °C.
Each purification fraction was mixed with an equal volume of 2×
protein loading buffer and boiled for 5 min. The samples were
separated on a 10% SDS-polyacrylamide gel then stained with
Coomassie blue to visualize. Purity was qualitatively quantified.
Thermal Shift Assay. Purified GGTase I complex was mixed with
compounds of interest in 1:40 ratio to final concentrations of 5 μM
and 100 μM in buffer A. This mixture was mixed with an equal
volume of 2× concentration of SYPRO Orange dye from Protein
Thermal Shift Dye Kit (Applied Biosystems) to the final volume of 10
μL in Hard-Shell 384-Well PCR Plate (Bio-Rad) and sealed with
Polyolefin Sealing Tape (Sarstedt). Fluorescence was observed using a
CFX384 Real-Time System (Bio-Rad) under the following con-
ditions; set to melt curve, step 1 (25 °C, 2 min) and step 2 (ramp to
98.6 °C, increasing 0.2 °C per 5 s cycle). Changes in fluorescence
over changes in temperature was calculated and plotted in Microsoft
Excel.
Geranylgeranylation Assay. The in vitro geranylgeranylation
assay was adapted from ref 52, 50 nM purified GGTase I complex, 20
μM GDP (Sigma-Aldrich), 10 μM NBD-FPP (Jena Bioscience), and
1 μM RhoA (Abcam) in buffer A to the final volume of 10 μL. The
reaction was incubated at 30 °C for 2 h. The reaction was stopped
with the addition of 10 μL of 2× protein loading buffer and boiled for
5 min. The samples were separated on a 10% SDS-polyacrylamide gel
and visualized with Typhoon FLA 9500 laser scanner with EGFP filter
(GE Healthcare). The gel was then stained with Coomassie blue to
visualize RhoA. EGFP and Coomassie blue intensities were quantified
using ImageJ following its background subtraction function.53 EGFP
intensities were calculated relative to the Coomassie blue intensity,
then relative EGFP intensities were calculated to the DMSO sample.
Authors
Sang Hu Kim − Department of Molecular Genetics, University of
Toronto, Toronto, Ontario M5G 1M1, Canada; orcid.org/
0000-0003-0597-0623
Chunliang Liu − Boragen, Inc., Durham, North Carolina
27709, United States
Yasheen Zhou − Boragen, Inc., Durham, North Carolina
27709, United States
Yong-Kang Zhang − Boragen, Inc., Durham, North Carolina
27709, United States
Cari McGregor − Boragen, Inc., Durham, North Carolina
27709, United States
Luke Steere − Boragen, Inc., Durham, North Carolina 27709,
United States
Brittany H. Frederick − Boragen, Inc., Durham, North
Carolina 27709, United States
C. Tony Liu − Boragen, Inc., Durham, North Carolina 27709,
United States
Luke Whitesell − Department of Molecular Genetics, University
of Toronto, Toronto, Ontario M5G 1M1, Canada
Complete contact information is available at:
https://pubs.acs.org/10.1021/acschembio.0c00290
Funding
S.H.K. is supported by a Mitacs Accelerate Fellowship
(IT13564) in association with Boragen Inc. L.E.C. is supported
by the Canadian Institutes of Health Research Foundation
Grant (FDN-154288). L.E.C. is a Canada Research Chair
(Tier 1) in Microbial Genomics & Infectious Disease and
codirector of the CIFAR Fungal Kingdom: Threats &
Opportunities program.
Notes
The authors declare the following competing financial
interest(s): L.E.C. and L.W. are co-founders and shareholders
in Bright Angel Therapeutics, a platform company for
development of novel antifungal therapeutics.
ACKNOWLEDGMENTS
We thank Cowen lab members for helpful discussions and the
Donnelly Sequencing Centre for whole genome sequencing.
■
REFERENCES
(1) Hawksworth, D. L., and Lu
■
ASSOCIATED CONTENT
■
̈
cking, R. (2017) Fungal Diversity
sı
* Supporting Information
The Supporting Information is available free of charge at
https://pubs.acs.org/doi/10.1021/acschembio.0c00290.
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the growth of efflux pump-compromised S. cerevisiae
strains, NMR spectra of benzoxaboroles 7, 8, and 9, and
reagents used in this study (PDF)
́
AUTHOR INFORMATION
■
Corresponding Author
Leah E. Cowen − Department of Molecular Genetics, University
of Toronto, Toronto, Ontario M5G 1M1, Canada;
orcid.org/0000-0001-5797-0110; Email: leah.cowen@
utoronto.ca
J
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