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Epigenetics
Epigenetics
Epigenetics
1559-2294
1559-2308
Taylor & Francis

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10.1080/15592294.2024.2400423
2400423
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Brief Report
Brief Report
Exploring fatty acids from royal jelly as a source of histone deacetylase inhibitors: from the hive to applications in human well-being and health
F. APARECIDA DOS SANTOS FRANCE ET AL.
EPIGENETICS
Aparecida dos Santos France Fernanda a
Maeda Debora Kazumi a
Rodrigues Ana Beatriz a
Ono Mai a
Lopes Nogueira Marchetti Franciele a
Marchetti Marcos Martins a
Faustino Martins Allana Cristina b
Gomes Roberto da Silva b
https://orcid.org/0000-0002-0285-1162
Rainho Cláudia Aparecida a
a Department of Chemical and Biological Sciences, Institute of Biosciences of Botucatu, São Paulo State University (UNESP) , Botucatu, SP, Brazil
b Department of Pharmaceutical Sciences, North Dakota State University , Fargo, ND, USA
CONTACT Cláudia Aparecida Rainho claudia.rainho@unesp.br Department of Chemical and Biological Sciences, Institute of Biosciences of Botucatu, São Paulo State University – UNESP, Rua Professor Doutor Antonio Celso Wagner Zanin, No. 250, Distrito de Rubiao, Junior, Botucatu, Sao Paulo18618-689, Brazil
10 9 2024
2024
10 9 2024
19 1 2400423Integra10 9 2024
Integra10 9 2024
05 5 2024
17 8 2024
30 8 2024
© 2024 The Author(s). Published by Informa UK Limited, trading as Taylor & Francis Group.
2024
The Author(s)
https://creativecommons.org/licenses/by-nc/4.0/ This is an Open Access article distributed under the terms of the Creative Commons Attribution-NonCommercial License (http://creativecommons.org/licenses/by-nc/4.0/), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited. The terms on which this article has been published allow the posting of the Accepted Manuscript in a repository by the author(s) or with their consent.

ABSTRACT

A differential diet with royal jelly (RJ) during early larval development in honeybees shapes the phenotype, which is probably mediated by epigenetic regulation of gene expression. Evidence indicates that small molecules in RJ can modulate gene expression in mammalian cells, such as the fatty acid 10-hydroxy-2-decenoic acid (10-HDA), previously associated with the inhibition of histone deacetylase enzymes (HDACs). Therefore, we combined computational (molecular docking simulations) and experimental approaches for the screening of potential HDAC inhibitors (HDACi) among 32 RJ-derived fatty acids. Biochemical assays and gene expression analyses (Reverse Transcriptase – quantitative Polymerase Chain Reaction) were performed to evaluate the functional effects of the major RJ fatty acids, 10-HDA and 10-HDAA (10-hydroxy-decanoic acid), in two human cancer cell lines (HCT116 and MDA-MB-231). The molecular docking simulations indicate that these fatty acids might interact with class I HDACs, specifically with the catalytic domain of human HDAC2, likewise well-known HDAC inhibitors (HDACi) such as SAHA (suberoylanilide hydroxamic acid) and TSA (Trichostatin A). In addition, the combined treatment with 10-HDA and 10-HDAA inhibits the activity of human nuclear HDACs and leads to a slight increase in the expression of HDAC-coding genes in cancer cells. Our findings indicate that royal jelly fatty acids collectively contribute to HDAC inhibition and that 10-HDA and 10-HDAA are weak HDACi that facilitate the acetylation of lysine residues of chromatin, triggering an increase in gene expression levels in cancer cells.

KEYWORDS

Human HDACs
10-HDA
10-HDAA
molecular docking
epi-drugs
epigenetic therapy
Fundação de Amparo à Pesquisa do Estado de São Paulo 10.13039/501100001807 07/59110-9 Conselho Nacional de Desenvolvimento Científico e Tecnológico 10.13039/501100003593 Coordenação de Aperfeiçoamento de Pessoal de Nível Superior 10.13039/501100002322 This study was supported by grants from Fundação de Amparo à Pesquisa do Estado de São Paulo [FAPESP grant# 07/59110-9], Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) and Coordenação de Aperfeiçoamento de Pessoal de Nível Superior - Brazil (CAPES) - Finance Code 001. FASF was granted a scholarship from the Brazilian Federal Agency for Support and Evaluation of Graduate Education (CAPES), within the scope of the Program CAPES-PrInt [process number 88887.310463/2018-00, mobility number 88887.570051/2020-00].
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pmcBackground

Epigenetic regulation orchestrates chromatin dynamics and gene expression and plays key roles in several biological processes. Biochemical chromatin modifications are part of a defined epigenetic programming to maintain transcriptional plasticity across multiple cell divisions during eukaryote development [1]. Epigenetic regulation includes complex mechanisms achieved through DNA methylation, typically in CpG dinucleotides, and histone post-translational modifications (PTMs). Both are biochemical marks of chromatin, heritable, reversible, and closely linked to the transcriptome. Moreover, environmental factors, lifestyle, and diet impact the epigenomic landscape and its function, allowing transcriptional adjustment to these external factors and conditions [2].

Honeybees are unique models for studying how diet determines two distinct phenotypes from one genome. Differential feeding during larval development with royal jelly (RJ) results in queens and workers exhibiting contrasting features, including differences in morphology, fertility, and longevity. Larvae destined to become queens receive a diet rich in royal jelly for an extended period, whereas those selected to become workers initially receive royal jelly and later transition to a diet consisting of honey and other compounds [3,4].

Royal jelly is a gelatinous product secreted by the hypopharyngeal and mandibular salivary glands of nurse honeybees. It comprises nutritional components such as carbohydrates, proteins, peptides, lipids, vitamins, and mineral salts. Although no single chemical compound in the RJ composition seems to be responsible for specifying larval fate, the caste differentiation process involves the integration of epigenetic and environmental factors [4]. In addition, natural products from the beehive, such as honey, propolis, and RJ, have been extensively used due to their health-promoting properties. They have attracted considerable interest as nutraceuticals and cosmeceuticals [5–7].

A recent study provided evidence of the effects of RJ on genome demethylation and expression of cancer-related genes in human cancer cell lines [8]. Spannhoff et al. suggested that the epigenetic regulation that controls the phenotypic plasticity of queens is, in part, mediated by the inhibitory activity of histone deacetylases (HDACs), by E-10-hydroxy-2-decenoic acid (10-HDA), a fatty acid exclusively found in RJ [9]. This evidence was recently reinforced by the discovery that honeybee histone proteins are extensively post-translationally modified, showing caste-specific signatures [10]. Together, these findings strengthen the hypothesis that chromatin modifications are key in establishing and maintaining caste-specific transcriptional programs in honeybees [11].

HDACs compose a family of enzymes highly conserved among species. Mammalian HDACs can be classified into four classes (I to IV), where classes I, II, and IV comprise the classical histone deacetylases, and class III consists of sirtuins, which differ in domain structure and enzymatic reaction. The human enzymes are well characterized and have known sequences and structural organization. HDACs 1, 2, 3, and 8 belong to class I, HDACs 4, 5, 7, and 9 belong to subclass IIa, HDACs 6 and 10 belong to subclass IIb, and HDAC11 belongs to class IV [12]. Although human HDACs have been extensively studied, little is known about their honeybee orthologs.

Interestingly, two independent research groups have reported the inhibitory effects of 10-HDA on recombinant human HDACs [9,13]. In addition to 10-HDA, RJ contains phenylbutyrate, a known HDAC inhibitor (HDACi) [14]. In this sense, we hypothesize that fatty acids from RJ could interact with human HDACs and inhibit their activity, thus being considered new HDACi candidates for epigenetic therapy. In this study, we searched for Apis mellifera classical histone deacetylase family members and described their similarities with human orthologs. Next, we performed in silico screening of 32 royal jelly-derived fatty acids with human HDACs, to predict the interactions of these chemical compounds with the deacetylase domain of human HDAC2 co-crystalized with SAHA (suberoylanilide hydroxamic acid), a widely studied HDACi, as a reference. Finally, the majoritarian fatty acids derived from RJ, i.e., 10-HDA and 10-HDAA (10-hydroxy-decanoic acid), were selected to validate their predicted inhibitory effects experimentally.

Materials and methods

Comparison between histone deacetylase orthologs in humans and honeybees

DNA sequences of histone deacetylase coding genes were retrieved from Apis mellifera (genome assembly Amel_4.5) and Homo sapiens (genome assembly GRCh38 - hg38 from Genome Reference Consortium), and their unique identity was confirmed on the National Center for Biotechnology Information (NCBI) records [15]. The amino acid sequences were retrieved from UniProt [16], and the domain information from InterProScan [17] and MOTIF (https://www.genome.jp/tools/motif) (Supplementary Table S1). The phylogram was built using Clustal Omega [18] default, which uses HMM (hidden Markov model) for the alignment engine and an enhanced version of mBed [19]. Then, it was visualized with iTOL [20]. Multiple protein sequence alignment was performed on Clustal Omega and visualized with Jalview 2.11.1.3 [21]. The pairwise comparison between Hs_HDAC1 and Am_Hdac1, Hs_HDAC2 and Am_Hdac1, Hs_HDAC3 and Am_Hdac3, and Hs_HDAC6 and Am_Hdac6 was calculated by Jalview 2.11.1.3 software.

Description of royal jelly-derived fatty acids

The list of fatty acids from royal jelly was obtained from previous reports [14,22–26] (Supplementary Table S2), and their chemical structures and identification numbers (CID) were manually curated from the PubChem database [27], which were used as input on ChemMine Tools Web Server [28]. Hierarchical clustering by structural similarities and physicochemical properties was calculated by OpenBabel Descriptors [29] using the single linkage method and Z-scores displayed. The distribution and intercorrelations of these chemical compounds were computed using the script from the literature, based on ”constellation plots” [30], to determine the cores and possible analog series of the fatty acids (Morgan fingerprint, minimum core size = 1). Complementary analysis of drug-like compounds was performed using the Platform for Unified Molecular Analysis (PUMA) [31], according to Lipinski’s rule of five.

Molecular docking simulations of interactions between fatty acids and human histone deacetylase catalytic domain

The molecular docking strategy was validated by self-docking (re-docking) [32–34], using a well-established HDAC inhibitor, suberoylanilide hydroxamic acid (SAHA/Vorinostat), co-located in the catalytic site, to reproduce the x-ray crystallography PDB ID: 4LXZ [35]. Root-Mean-Square Deviation (RMSD) was calculated using the LigRMSD web server [36] between the crystallized ligand and the docked ligand, demonstrating high similarity (0.84 Å). The tested cavities were the allosteric site of activation by inositol phosphates and the catalytic site of class I HDACs. The SWISS-MODEL web server was used to model suitable templates for docking simulations [37]. For the allosteric and catalytic sites, the chosen models were HDAC3 bound to inositol tetraphosphate (PDB ID: 4A69) [38] and HDAC2 (PDB ID: 4LXZ) [35], respectively. In silico screening was performed using AutoDock Vina [39] through the PyRx graphical interface [40]. HDAC3 coordinates: X = 24.038 Å, Y = 48.472 Å and Z = 21.341 Å, sizes X, Y and Z = 15 Å. HDAC2 coordinates: center X = 24.9537 Å, Y = −16.1707 Å and Z = 1.9851 Å, sizes X = 13.9479 Å, Y = 11.4023 Å and Z = 13.0559 Å. Exhaustiveness = 10. Target preparations were conducted using BIOVIA Discovery Studio 2020 (Dassault Systèmes, San Diego, CA, USA). The search area was delimited in Chain A. Ligand structures derived from PubChem data were converted using OpenBabel 2.3.2 software [29], and all geometries were virtually constructed and optimized in molecular mechanics (MM+) and semiempirical Austin Model 1 (AM1) methods, using HyperChem 8.0.5 software [41] for geometry optimization. The putative interactions with the residues were visualized on the Protein-Ligand Interaction Profiler (PLIP) [42].

Cell-free and in vitro HDAC activity assays

Two major RJ fatty acids were selected for experimental validation: 10-hydroxy-2-decenoic acid (10-HDA, CID 5312738; ChemCruz, Santa Cruz Biotechnology, TX, USA) and 10-hydroxydecanoic acid (10-HDAA; CID 74300; Merck, Sigma-Aldrich, MO, USA). Both were diluted in dimethyl sulfoxide (DMSO; Sigma-Aldrich, MO, USA) to a stock solution at 1 M. The cell-free Histone Deacetylase Assay Kit, Fluorometric (Merck, Sigma-Aldrich, MO, USA) was performed according to the manufacturer’s instructions for HDAC inhibition testing, using 1.25 µM of TSA and 1.25 µM of SAHA (Merck, Sigma-Aldrich, MO, USA) as positive controls (Supplementary Figure S1). 10-HDA and 10-HDAA, alone or in combination, were tested at 2.5, 5, and 10 mM. Fluorescence was read at 365/415-445 nm using the GloMax® Explorer Multimode Microplate Reader (Promega, Madison, WI, USA).

In vitro assay was performed in three independent experiments after exposing the breast cancer MDA-MB-231 cell line to 100 μM of 10-HDA and 10-HDAA, alone or in combination, and 500 nM of TSA, for 12 h and 24 h. Post-treatment, cells were processed for nuclei isolation using the NuCLEAR™ Extraction Kit (Merck, Sigma-Aldrich, MO, USA), according to the manufacturer’s instructions. The nuclear extracts were used for the Histone Deacetylase 3 (HDAC3) Activity Assay Kit (Merck, Sigma-Aldrich, MO, USA), following the manufacturer’s instructions. Relative HDAC activity was calculated using untreated control values as 100% HDAC activity.

In vitro effects of 10-HDA and 10-HDAA in the expression of HDAC genes in HCT116 and MDA-MB-231 cell lines

Cell culture and in vitro treatments

HCT116 and MDA-MB-231 cell lines were cultured according to a previous study of our research group [43]. Cells were treated in triplicates for 72 hours, with 10-HDA (ChemCruz, Santa Cruz Biotechnology, TX, USA), 10-HDAA (Merck, Sigma-Aldrich, MO, USA), or both drugs, at a concentration of 100 μM, using dimethyl sulfoxide (DMSO; Merck, Sigma-Aldrich, MO, USA) as control. Following the treatment, cellular pellets were collected and stored at −80°C.

Total RNA extraction and cDNA synthesis

Total RNA was extracted using the AllPrep DNA/RNA/miRNA Universal Kit (Qiagen, Hilden, Germany) and purified with DNAse I amplification grade (Invitrogen, Carlsbad, CA, USA). The complementary DNA (cDNA) was synthesized with random primers using SuperScript™ II Reverse Transcriptase (Invitrogen, Carlsbad, CA, USA), according to the manufacturer’s instructions.

Gene expression analysis by Reverse Transcriptase – quantitative Polymerase Chain Reaction (RT-qPCR)

RT-qPCR was performed using GoTaq® qPCR Master Mix 2X (Promega, Madison, WI, USA), 100 nM forward and reverse primers, and 250 ng cDNA in a final volume of 10 µL. The reactions were performed in triplicate, using the StepOne™ Real-Time PCR System (Applied Biosystems, Foster City, CA, USA), with the following conditions: 1 cycle at 95°C for 10 minutes, 40 cycles of denaturation at 95°C for 30 seconds, followed by annealing/extension at 60°C for 1 minute. Gene expression was calculated based on 2−ΔΔCt method, normalized with Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) expression levels. The primer sequences are described in Supplementary Table S3.

Statistical analysis

Statistical analyses were performed using GraphPad Prism Version 10.2.2 (GraphPad Software, Boston, MA, USA), based on the ANOVA test and Dunn’s correction for multiple comparisons, to determine statistical significance (p < 0.05) of the experimental data compared with untreated controls.

Results

Histone deacetylase domains are conserved in Apis mellifera and Homo sapiens classical HDACs

Four members of the histone deacetylase family were found in Apis mellifera genome annotation: Am_Hdac1, Am_Hdac3, Am_Hdac5, and Am_Hdac6 (Supplementary Table S1, Supplementary Figure S2A), which are highly conserved (Supplementary Figure S2B) and have domain similarities to human HDACs (Supplementary Figure S2C and S2D). Histidine, tyrosine and aspartate residues were remarkably conserved in the catalytic site (Supplementary Figure S2C). The amino acid sequences of Hs_HDAC1 and Am_Hdac1 showed 76.18% identity and presented the highest alignment score (20130.0) among all orthologs. While Hs_HDAC2 and Am_Hdac1 also showed high identity (76.02% and 20120.0 scores), the alignment of class II enzymes resulted in lower identity levels (40.77% for Hs_HDAC5 and Am_Hdac5 comparison, and 36.16% for Hs_HDAC6 and Am_Hdac6). Based on these data, Hs_HDAC1 and Hs_HDAC2 were selected as the best representative receptors for molecular docking simulations between fatty acids as ligands and the deacetylase domain.

Royal jelly-derived fatty acids have drug-like properties

The chemical structures of the 32 fatty acids are illustrated in Supplementary Figure S3. These compounds were classified into two distinct groups. The first includes seven short-chain fatty acids (SCFA) showing a terminal ring associated with a cyclic ester (lactone), sharing the aromatic ring as the main feature, as well as cyclic esters in delta-decalactone and 6-propyloxan-2-one. In addition, 25 fatty acids present a mid-chain length (MCFA) that is generally 8–12 carbon atoms long and share similarities in their physicochemical properties (Supplementary Figure S4A and S4B). Most fatty acids passed Lipinski’s rule of five, except 10-acetyloxy decanoic acid, (11S)12-dihydroxydodecanoic acid, and 3-hydroxydodecanedioic acid (Supplementary Table S4). Although they represent different analog series, there was no correlation with the predicted energy values, and they were grouped into a few cores of the analog series (Supplementary Figure S5).

Royal jelly-derived fatty acids might interact with the human HDAC2

We used two well-known HDAC inhibitors, SAHA and TSA, and the proposed catalytic deacetylation mechanism (Figure 1a) as references to evaluate the possible binding model of selected fatty acids to the active site of Hs_HDAC2 based on in silico docking simulations. Similar to SAHA and TSA, docking solutions indicate that 10-HDA and 10-HDAA may interact with the Hs_HDAC2 deacetylase domain through hydrogen bonds with Tyr308 amino acid residues and hydrophobic interactions with Phe155 and Phe210 residues. Interestingly, these compounds can also form salt bridges with Hist145, His146, and His183 (Figures 1b–e). Figure 1. a) simplified illustration of the HDAC catalytic deacetylation mechanism upon the acetylated lysine residue at the active site. The zinc ion is coordinated by two tandem histidine residues, which make hydrogen bonds to two aspartic acids. One of these histidines serves as a base for the nucleophilic attack at the substrate carbonyl by a water molecule hydrolysis. The tetrahedral oxyanion intermediate arising from nucleophilic attack by water is illustrated in the center following the acetate liberation from the N-terminal histone tail (right). (b-e) on the left 2D molecular structures of the tested compounds; in the center, the molecular docking prediction of the lowest energy poses of well-known HDACi (hydroxamic acids SAHA and TSA) and the carboxylic acids derived from RJ (10-HDA and 10-HDAA); on the right side, a simplified diagram of the predicted interactions and their respective distances. The HDAC2 protein is shown in a gray ribbon. The active site residues are shown in ball-and-stick style, with hydrogen in white, nitrogen in blue, oxygen in red, and zinc cation in pink. The ligands are shown in light blue, and the 10-HDAA is highlighted in light green to distinguish it from the 10-HDA. The 2D molecular structure and the predicted interactions represent hydrogen bonds in blue dotted lines, zinc interactions with black dashed lines, and salt bridges in yellow dotted lines with a charge center indicated.

The binding affinities for the best docking solutions of the fatty acids analyzed are presented in Table 1. Among the seven SCFAs, the binding energy values ranged from −5.5 kcal/mol (6-propyl hexan-2-one) to −7.3 kcal/mol for dihydroferulic acid (DHFA), a chemical compound that has never been associated with HDAC inhibition. Phenylbutyrate, known to inhibit HDACs, presented an unfavorable energy value of −5.8 kcal/mol. Interestingly, 75% of MCFAs (19 out of 25 chemical compounds) presented similar binding energy values, ranging from −6.1 to −6.7 kcal/mol. The most abundant MCFAs were selected for experimental validation, corresponding to 60–80% of total organic acids in royal jelly, 10-HDA, and 10-HDAA.Table 1. Docking scores for the lowest energy poses of fatty acids derived from royal jelly and deacetylase domain of Hs_HDAC2 obtained from AutoDock Vina.

Rank	CID	Compound Name	Energy (kcal/mol)*	
Short-chain fatty acids	
1	14340	Dihydroferulic acid (DHFA)	−7.3	
2	12810	Delta-decalactone	−6.0	
3	7456	Methylparaben	−6.3	
4	7469	4’-Hydroxyacetophenone (Piceol)	−6.3	
5	2214	Acetovanillone (Apocynin)	−6.3	
6	20354	Phenylbutyrate	−5.8	
7	12777	6-Propyloxan-2-one	−5.5	
Mid-chain fatty acids	
1	11637116	(S)-11-Hydroxydodecanoic acid	−6.7	
2	26612	3-hydroxydecanoic acid	−6.7	
3	91394357	(10 R,11 R)-10,11-Dihydroxydodecaneoic acid	−6.7	
4	11564887	(11S)-11,12-Dihydroxydodecanoic acid	−6.6	
5	1713086	9-oxo-2-decenoic acid	−6.6	
6	24990605	3,9-dihydroxydecanoic acid	−6.6	
7	11535975	3,11-dihydroxydodecanoic acid	−6.5	
8	547024	11-Oxododecanoic acid	−6.5	
9	9859090	3,10-dihydroxydecanoic acid	−6.5	
10	16663321	3-hydroxydodecanedioic acid	−6.4	
11	86301641	(E)-10-Acetoxy-2-decenoic acid	−6.4	
12	13497206	10-acetyloxydecanoic acid	−6.3	
13	25141159	(E)-9,10-Dihydroxy-2-decenoic acid	−6.3	
14	5192	Sebacic acid	−6.3	
15	5312738	(E)-10-hydroxydec-2-enoic acid (10-HDA)	−6.3	
16	5312739	(E)-9-Hydroxy-2-decenoic acid	−6.3	
17	6079438	2-Octenedioic acid	−6.3	
18	181528	(E)-2-Decene-1,10-dioic acid	−6.2	
19	379	Caprylic acid	−6.1	
20	167627	7-Hydroxyoctanoic acid	−5.9	
21	4575239	9-hydroxydecanoic acid	−5.8	
22	537408	9-hydroxynonan-2-one	−5.8	
23	69820	8-Hydroxyoctanoic acid	−5.8	
24	74300	10-Hydroxydecanoic acid (10-HDAA)	−5.8	
25	521747	Methyl 3-hydroxydecanoate	−5.4	
*Binding affinity energy calculated by AutoDock Vina.

10-HDA and 10-HDAA inhibit human HDACs

The inhibitory activity of the 10-HDA and 10-HDAA was tested in a commercially available cell-free nuclear extract of human HeLa cells. Although less effective than SAHA and TSA, 10-HDA and 10-HDAA reduced HDAC activity, alone or in combination (Figure 2a). In addition, after in vitro exposure of the cell line derived from breast cancer, MDA-MB-231, it was observed that after 12 h of exposure, both compounds, alone or in combination, inhibited HDAC3 activity, like TSA (p ≤0.05). Although no effect was observed after 24 h of exposure to 10-HDA or 10-HDAA, an increase in the deacetylase activity was detected when both compounds were associated (p ≤0.001) (Figure 2b). Figure 2. a) relative HDAC inhibition using 2.5 mM, 5 mM, and 10 mM of 10-HDA and 10-HDAA, alone or in combination, compared to each internal control in a direct cell-free activity assay. b) HDAC3 inhibition assay after in vitro exposure of MDA-MB-231 cells to 10-HDA and/or 10-HDAA (100 μM each), SAHA and TSA (500 nM) during 12 and 24 h. c and d) expression levels of class I HDACs encoding genes after 72 h of in vitro exposure to 10-HDA and/or 10-HDAA (100 μM each). (* = p ≤ 0.05, ** p ≤ 0.01, and *** = p ≤ 0.001).

10-HDA and 10-HDAA modulate the expression levels of human class I HDAC genes

To investigate whether inhibiting enzyme activities was subjacent to the downregulation of gene transcription, we determined the expression levels of four nuclear class I HDACs in two human cancer cell lines (HCT116 and MDA-MB-231). The exposure to 100 μM of 10-HDA and/or 10-HDAA showed a discrete increase in the expression level of HDAC1 in HCT116 cells (Figure 2c). Furthermore, an increase in the expression levels of the HDAC3 gene was detected in MDA-MB-231 cells, especially when both fatty acids were combined (Figure 2d).

Discussion

HDACs are proteins involved in chromatin remodeling by removing acetyl groups in the amino-terminal tail of histones, controlling the levels of histone acetylation and gene expression [12]. Human HDACs have been implicated in the pathogenesis of cancer, diabetes, and immune and neurological diseases [44–46]. Currently, pan- and selective-HDAC inhibitors, HDAC-based dual-target inhibitors, and HDAC degraders based on proteolysis targeting chimeras (PROTACs) are alternative strategies to target this class of enzymes in epigenetic therapy protocols [47].

The chemical composition of RJ has been studied over the last 80 years [48,49]. Since then, hydroxy-fatty acids have been recognized as the major component of the lipid portion of this natural compound [25]. In particular, 10-HDA, a fatty acid exclusively found in RJ composition, has been associated with inhibitory histone deacetylase activity [9]. First, 10-HDA has attracted attention because of its antibacterial [50] and antitumor [51,52] properties. The health-promoting activities of 10-HDA have been extensively explored in different age-related human disorders, including neurodegenerative and cardiovascular diseases and cancer [53]. In this study, we investigated the potential of fatty acids derived from RJ as new HDAC inhibitor candidates.

In summary, we have shown that human and honeybee HDACs are conserved and that orthologs bring together amino acid residues involved in the catalytic mechanism. Predicted interactions between the catalytic domain of Hs_HDAC2 and fatty acids from RJ based on molecular docking simulations were validated using a pan-HDAC inhibition assay, suggesting that both species share conserved catalytic cores and probably similar catalytic mechanisms. In vitro treatment of human cancer cell lines with the two major fatty acids (10-HDA and 10-HDAA) revealed that these compounds can inhibit the intrinsic nuclear activity of HDACs after 12 h of exposure. However, this effect was lost after 24 h.

Although the literature lacks details about HDAC’s evolutionary origins, they are believed to exist in archaea, prokaryotes, and eukaryotes [54,55]. HDAC family members targeting histone or non-histone proteins preceded the evolution of histone proteins [56] and are an important interface between environmental changes and the chromatin landscape [57]. For instance, HDACs play a key role in phenotypic plasticity in Apis mellifera during larval development [3,4]. In general, histone sequences and the chromatin-modifying machinery are conserved. For example, PTMs on the histone tails of histones H3.1, H3.3, and H4 in Apis mellifera have been characterized [10], and histone methyltransferases and demethylases have been described [11]. In addition to the modulation of caste switching [58], the involvement of histone acetylation in aversive memory [59] and behavior has been described in honeybees [60]. A more recently published article indicated that 10-HDA, zinc, and potassium concentrations are strongly correlated with caste differentiation, among other control mechanisms [13], such as microRNAs [61–63] and methionine concentration [64].

Although no Apis mellifera HDAC crystal structures exist in public databases, histone deacetylases encoded in the honeybee genome are similar to human class I and class II enzymes [9]. If 10-HDA inhibits class I and class II HDACs in honeybees and the recombinant HDACs tested by Spannhof et al., it is reasonable to hypothesize that 10-HDA could similarly inhibit human HDACs. Our molecular screening and biochemical assay data provide evidence that fatty acids from RJ can interact with the deacetylase domain and inhibit the activity of class I human HDACs. However, it is lower than the inhibition by well-known HDACi. The chemical structures of SAHA and TSA share a common pharmacophore feature, including a capping group (aromatic ring) binding on the surface of the active site pocket and a zinc-binding group (ZBG), with a hydroxamic acid group that chelates the zinc ion in the catalytic pocket [12]. Although hydroxamic acid is thought to be critical for inhibition potency, there are small molecules recognized as HDACi, such as butyric acid, which is detected in RJ, and valproic acid, that lack the hydroxamic acid functional group [65]. In this context, the molecular docking predictions indicate that the short-chain fatty acids from RJ present limited contact with the catalytic pocket and are probably associated with weak inhibitory activity. In addition, MCFA derived from RJ lacks the hydroxamic group to bind to zinc, which is related to its weak inhibitory capability despite its better position in the tubular structure with a zinc atom at its base, which is necessary for zinc-dependent deacetylation.

Interestingly, RJ has shown bioactive effects in in vitro models using human breast carcinoma cells [66–68] and in a recently published clinical trial of renal cell carcinoma [69]. Few clinical trials are ongoing to explore the potential use of RJ in cancer patients submitted to radiotherapy (for example, ClinicalTrials.gov - NCT01871155 and UMIN-CTR - UMIN000020152) or other diseases (ClinicalTrials.gov - NCT06288204). Although RJ is not a direct focus of drug discovery studies, distinct chemical compounds in its composition have been investigated to assess their potential therapeutic benefits. Further studies are needed to validate and advance our understanding of the potential applications in drug development. No specific chemical compounds derived from RJ have been approved by the US Food and Drug Administration (FDA) for therapeutic or medicinal use. Nevertheless, SCFA phenylbutyrate and its derivatives have been identified as HDACi in preclinical and clinical trials for cancer, diabetes, and neurological disorders [70]. For example, the carboxylic acid pivaloyloxymethyl butyrate (Pivanex; AN-9) has been explored as an HDAC inhibitor in leukaemic cells and solid tumours, including lung, breast, and colorectal carcinomas [71].

Based on the molecular docking results indicating similar binding energy values for the MCFA group, the major fatty acids of royal jelly (the 10-HDA and 10-HDAA) were selected to test the in vitro inhibition of HDAC activity, in HCT116 and MDA-MB-231 cell lines. These compounds, isolated or in combination, altered the gene expression levels of HDAC1 in the HCT116 cell line and HDAC3 in MDA-MB-231, both of which encode nuclear zinc-dependent histone deacetylases (class I HDACs). Moreover, our findings indicate that 10-HDA and 10-HDAA, whether alone or in combination, interact with the catalytic tunnel of nuclear histone deacetylases, inhibiting their enzymatic activity. This effect, observed in the HDAC3 assay after 12 h of exposure, is similar to that of TSA, an hydroxamic acid. After 24 h of exposure, no effects were observed in the samples treated with the compounds alone. However, a significant increase in HDAC3 activity was observed in cells treated with 10-HDA and 10-HDAA in combination. These findings suggest that, although the biochemical inhibition is ceased, there are lasting effects in chromatin modulation, as a consequence of HDAC inhibition. To better evaluate these results, we explored the RNA-seq data available on GEO Datasets (GSE236280) of MDA-MB-231 exposed to a known histone deacetylase inhibitor panobinostat (data not shown) [72]. In this study, differentially expressed genes (DEGs) showed an increase in expression in response to panobinostat treatment, compared with the untreated control. In addition, HDAC3 was detected among these upregulated DEGs. These findings reinforce our results and corroborate the hypothesis that 10-HDA and 10-HDAA are HDAC inhibitors that favor the acetylation of lysine residues of chromatin, triggering an increase in global gene expression levels. Alternatively, indirect effects of fatty acid metabolism and changes in the levels of acetyl radicals could change chromatin acetylation levels and interfere with the equilibrium between the opposite activities of histone acetyltransferases (HATs) and HDACs. Although the subjacent mechanisms are not completely understood, HDAC3 is a nuclear and cytoplasmic enzyme that has recently been functionally linked to the Nuclear Receptor Co-Repressor 1 (NCoR1) in regulating energy metabolism and glucose homeostasis. Further studies are needed to advance the understanding of how HDAC3 and the co-repressive complexes NCoR1/2 modulate metabolic processes and are implicated in human diseases [73]. Our study advances the knowledge regarding the effects of 10-HDA and 10-HDAA in inhibiting human HDACs. Moreover, other RJ-derived fatty acids could be explored in further studies regarding their potential for discovering new HDACi, considering that the investigated fatty acids presented similar binding energy values.

Interestingly, a previous study reported an increase in Am_Hdac3 expression in bee larvae exposed for long periods to 10-HDA [74], highlighting this change in expression as a late effect of HDAC inhibition, which results in increased acetylation of histone lysine residues. In turn, it triggers the relaxation of the chromatin structure, culminating in an increase in gene expression levels, including those encoding HDACs.

Conclusion

Several studies have explored the biological and pharmacological properties of bee products, but few have sought to unravel the mechanisms of action and targets of their bioactive molecules. The remaining gap concerns specificity and functional impacts on human health. This research aimed to advance our understanding of bioactive bee products by focusing on the role of fatty acids from royal jelly in epigenetic regulation and identifying opportunities for nutritional epigenetics and epi-drug discovery studies. In this context, herein, we demonstrate that despite the slight heterogeneity of royal jelly-derived fatty acids, they are predicted to inhibit histone deacetylases given their similar binding energies as predicted by molecular docking simulations. Notwithstanding, in the biological context, the major fatty acids that are available at higher concentrations, 10-HDA and 10-HDAA, alone or combined, inhibit human HDACs similarly as well-established hydroxamic acids for up to 12 h in the MDA-MB-231 breast cancer cell line. This direct inhibitory effect on enzyme activity is rapidly lost but there are lasting effects on gene expression upregulation of target genes. Broadly, as new epi-drugs are a promising area in chemoprevention and therapeutic strategies for complex human diseases, our data point to the potential of widely used natural products as weak inhibitors of HDACs.

Supplementary Material

Supplemental Material

-) Supplementary material _Tables_France et al_May2024.docx

Disclosure statement

No potential conflict of interest was reported by the author(s).

Author’s contributions

FASF, RSG, and CAR conceived and designed the approach, interpreted the results, wrote, and revised the manuscript. FASF, DKM, ABR, and MO conducted the experiments. FASF, FLNM, MMM, and ACFM conducted bioinformatic analysis and data interpretation. All authors read and approved the final version of the manuscript.

Data availability statement

All data generated or analyzed during this study are included in this published article and its supplementary files.

Supplementary material

Supplemental data for this article can be accessed online at https://doi.org/10.1080/15592294.2024.2400423
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