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. 2018 Mar 25;233(9):7143–7156. doi: 10.1002/jcp.26537

Omega‐3 fatty acids modulate the lipid profile, membrane architecture, and gene expression of leiomyoma cells

Md Soriful Islam 1,2, Clara Castellucci 3, Rosamaria Fiorini 4, Stefania Greco 1, Riccardo Gagliardi 5, Alessandro Zannotti 1, Stefano R Giannubilo 6, Andrea Ciavattini 6, Natale G Frega 7, Deborah Pacetti 7,✉, Pasquapina Ciarmela 1,8,✉
PMCID: PMC13482449  PMID: 29574773

Abstract

Uterine leiomyomas (fibroids or myomas) are the most common benign tumors of premenopausal women and new medical treatments are needed. This study aimed to determine the effects of omega‐3 fatty acids on the lipid profile, membrane architecture and gene expression patterns of extracellular matrix components (collagen1A1, fibronectin, versican, or activin A), mechanical signaling (integrin β1, FAK, and AKAP13), sterol regulatory molecules (ABCG1, ABCA1, CAV1, and SREBF2), and mitochondrial enzyme (CYP11A1) in myometrial and leiomyoma cells. Myometrial tissues had a higher amount of arachidonic acid than leiomyoma tissues while leiomyoma tissues had a higher level of linoleic acid than myometrial tissues. Treatment of primary myometrial and leiomyoma cells with eicosapentaenoic acid (EPA) or docosahexaenoic acid (DHA) reduced the monounsaturated fatty acid (MUFA) content and increased the polyunsaturated fatty acid (PUFA) content in both cell types. Myometrial and leiomyoma cell membranes were in the liquid‐crystalline phase, but EPA‐ and DHA‐treated cells had decreased membrane fluidity. While we found no changes in the mRNA expression of ECM components, EPA and DHA treatment reduced levels of ABCG1, ABCA1, and AKAP13 in both cell types. EPA and DHA also reduced FAK and CYP11A1 expression in myometrial cells. The ability of omega‐3 fatty acids to remodel membrane architecture and downregulate the expression of genes involved in mechanical signaling and lipid accumulation in leiomyoma cells offers to further investigate this compound as preventive and/or therapeutic option.

Keywords: ABCG1 and ABCA1, lipid profile, mechanical signaling, omega‐3 fatty acids, uterine leiomyoma

Myometrial and leiomyoma cell membranes were in the liquid‐crystalline phase, but EPA‐ and DHA‐treated cells had decreased membrane fluidity. EPA and DHA treatment reduced levels of ABCG1, ABCA1, and AKAP13 in both cell types. EPA and DHA also reduced FAK and CYP11A1 expression in myometrial cells. The ability of omega‐3 fatty acids to remodel membrane architecture and downregulate the expression of genes involved in mechanical signaling and lipid accumulation in leiomyoma cells offers to further investigate this compound as preventive and/or therapeutic option.

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1. INTRODUCTION

Uterine leiomyomas (fibroids or myomas), originating from myometrial smooth muscle cells of the uterus, are the most common benign tumors of fertile women (Bulun, 2013; Protic et al., 2016). Uterine leiomyomas affect about 77% of women of reproductive‐age and approximately 25% of reproductive‐age women bear clinically apparent tumors (Buttram & Reiter, 1981; Cramer & Patel, 1990). The incidence and severity of symptoms typically depend on the size, number, and location of the fibroids (Buttram & Reiter, 1981). The common symptoms associated with uterine leiomyomas are irregular and excessive menstrual bleeding, which often causes anaemia, pain in the back of the legs, pelvic pain or pressure, bowel and bladder dysfunctions, pressure sensation in the lower abdomen, pain during sexual intercourse, infertility, and recurrent abortion (Buttram & Reiter, 1981; Evans & Brunsell, 2007; Marsh & Bulun, 2006; Stewart, 2001). Uterine leiomyomas are currently the most common indication for hysterectomy in the world. Despite their high prevalence, significant associated health problems and huge economic impact on the healthcare system, relatively little is understood about the etiology and pathophysiology of uterine leiomyomas and effective therapeutic strategies are lacking. The prevalence of uterine leiomyoma is over three‐fold higher in black women compared to white women (Marshall et al., 1997). In addition, age (late reproductive years), heredity, nulliparity, obesity, polycystic ovary syndrome, diabetes, hypertension, oral contraceptives, and hormone replacement therapy are major risk factors associated with this tumor (Flake, Andersen, & Dixon, 2003; Okolo, 2008; Walker & Stewart, 2005).

To date, genetic and epigenetic factors, sex steroids, growth factors, cytokines, chemokines, inflammation, and extracellular matrix (ECM) components are known factors involved in the pathogenesis of leiomyoma (Ciarmela et al., 2011; Gallagher & Morton, 2016; Islam, Akhtar, Segars, Castellucci, & Ciarmela, 2017; Islam et al., 2013; Protic et al., 2016; Yang, Mas, Diamond, & Al‐Hendy, 2016). ECM components, mainly collagen1A1, fibronectin and versican are overexpressed in leiomyoma (Malik & Catherino, 2012), and their upregulation is induced by activin A (Islam et al., 2014). ECM proteins transmit mechanical signals from the outside of the cell to the cell interior through transmembrane integrin proteins. The functional activity of integrins may be modulated by modification of the fluidity of the cell membrane caused by alterations of lipid membrane composition.

Lipid homeostasis is fundamental for cellular function because of the key role of lipids in cell signaling, energy storage, and membrane functions, which are strictly dependent on membrane fluidity (Ibarguren, López, & Escribá, 2014). Lipidomics, or the comprehensive and quantitative analysis of lipid components in a given system, has been used to describe many pathological processes (Han, 2005; Rasmiena, Ng, & Meikle, 2013; Santos & Schulze, 2012). However, to our knowledge it has never been used to investigate uterine leiomyoma. Omega‐3 fatty acids (also called ω3 PUFA or n − 3 PUFA) are polyunsaturated fatty acids (PUFA) that are referred to as ‘essential fatty acids’ because they cannot be synthesized by mammals and must be obtained from the diet (Rose & Connolly, 1999). ω3 PUFA are commonly found in marine and plant oils and are being increasingly promoted as important dietary components for health and disease prevention. The active components of fish oil are generally considered to be the two ω3 PUFA, eicosapentaenoic acid (EPA), and docosahexaenoic acid (DHA) (Whelan & Rust, 2006). Data from experimental and clinical studies have provided evidence that n‐3 fatty acids are anti‐inflammatory and immunomodulatory, making n‐3 fatty acids potential therapeutic agents for inflammatory and autoimmune diseases (Calder, 2001).

The aim of the present study was to evaluate: (i) if the fatty acid profile of leiomyoma differs from that of healthy myometrium; (ii) if ω3 fatty acids modulate the lipidomic profile of the cell; and (iii) if ω3 fatty acids modulate the expression of ECM and mechanical signaling components as well as intracellular proteins such us sterol regulatory molecules involved in steroid synthesis and transport of cholesterol.

2. MATERIALS AND METHODS

2.1. Drugs and chemicals

EPA (cis‐5,8,11,14,17‐Eicosapentaenoic acid) and DHA (cis‐4,7,10,13,16,19‐Docosahexaenoic acid) were purchased from Sigma–Aldrich (St. Louis, MO). Both compounds were dissolved in ethanol (absolute) at 30 mM and then further diluted with culture medium to reach 50 μM at the time of cell treatment.Water (H2O) of Liquid Chromatography‐Mass Spectrometry (LC‐MS) grade and purchased from Sigma–Aldrich. All others solvents and reagents were of analytical grade. Supelco 37 Component FAME Mix (certified reference material grade) was employed as standard mixture and purchased from Supelco (Bellefonte, PA).

2.2. Leiomyoma and myometrial tissue collection

The study included premenopausal Caucasian women (41–49 years old) with symptomatic fibroids. Women did not receive hormonal treatment during the 3 months before to surgery. All patients gave their informed consent and the permission of the Human Investigation Committee was granted. Fibroid and myometrial tissues were obtained by hysterectomy or laparotomic myomectomy. The location of fibroids was submucosal and intramural, and their size range was 3–10 cm of diameter. Fibroid tissue was defined based on well‐established histopathologic criteria.

2.3. Primary cell cultures

Myometrial and leiomyoma samples were collected in Hanks'Balanced Salt Solution (HBSS) (Euroclone, Milan, Italy) at the time of surgery. Tissues were washed twice with Dulbecco's PBS (Invitrogen, Life Technologies, Carlsbad, CA) to remove excess blood. Samples were cut into small pieces with 0.1% collagenase type 8 (Serva Electrophoresis GmbH, Heidelberg, Germany) solutions in serum‐free Dulbecco's Modified Eagle Medium (DMEM) (Sigma–Aldrich) containing 1% penicillin‐streptomycin (Euroclone), 50 μg/L gentamicin (Lonza, Walkersville, MD), and 1% Amphotericin B (Lonza). Tissues were then incubated at 37 °C for about 3–5 hr in a water bath with manual shaken until completely digested. Digested cell suspensions were centrifuged at 1,200 rpm for 10 min, and washed once with fetal bovine serum (FBS) (Sigma–Aldrich). Finally, the cell pellet was dispersed in DMEM containing 10% FBS, 1% penicillin–streptomycin (Euroclone), 50 mg/L gentamicin (Lonza), and 1% Amphotericin B (Lonza), and plated in T25 or T75 plastic dishes, and maintained with same media at 37 °C in 95% air‐5% CO2. The growth medium was changed after 48 hr or 72 hr to remove unattached cells and then subsequently twice a week. The purity of cells was assessed by immunocytochemical staining with specific smooth muscle cells marker, monoclonal mouse anti‐α‐smooth muscle actin (α‐SMA) (Sigma–Aldrich). The lower passage number (up to 5) of cells was used for experiments to avoid changes in phenotype and gene expression.

2.4. Transmethylation of uterine myometrium and uterine fibroma tissue fatty acids

The direct transmethylation reaction of the tissue fatty acids (FA) was performed adapting the method of Harris (Harris, Pottala, Vasan, Larson, & Robins, 2012). For each sample, 80 mg of tissue were transferred and freeze dried in a 2 ml screwed cap vial. A total of 250 μl of a solution of BF3 (12% in MeOH anhydrous from Sigma–Aldrich®) and 250 μl of hexane were added in sequence to the freeze‐dried sample. The vial was capped and the suspension was vortexed for 15 s. Then, the vial was put in the oven at 100 °C for 15 min. After cooling, 250 μl of H2O were added to the sample and, after recapping the vial, the mixture was vortexed for 20 s. Two phases were obtained after centrifugation. A total of 150 μl of the upper hexane phase, containing the fatty acids methyl esters (FAME), were transferred in a 300 μl glass insert in a screwed cap vial. The solution was dried under nitrogen flow, and the FAME were dissolved in 50 μl of hexane. Samples were stored at −20 °C till the gaschromatographic (GC) analysis.

2.5. Transmethylation of cell pellet fatty acids

Transmethylation reaction was performed modifying the method described by Harris (Harris et al., 2012). Briefly, 30 mg of defrosted cell pellets were suspended in 300 μl of water. The suspension was transferred to a 2 ml screwed cap vial and freeze dried. The transmethylation method to obtain the FAME from the freeze‐dried cell pellets was the same employed for the derivatization of the FA from the freeze dried uterine myometrium and fibroma tissues reported above. Samples were stored at −20 °C till the GC analysis.

2.6. GC‐FID analysis of FAME

GC analysis of the FAME in cell pellets and uterine tissues samples was performed with a Varian® 430 GC. The chromatographic system was equipped with a split/splitless injector, a 100 m Varian® capillary column Select™ FAME (0.25 mm, 0.25 μm, #CP7421) and a flame ionization detector (FID). A total of 0.3 μl of sample were injected with a split ratio of 2. The injector and the FID were maintained at 260 °C. Helium (He) was employed as carrier gas with a flow rate of 1.6 ml/min. A gradient of temperature was set, with the oven starting at 160 °C and reaching 240 °C in 20 min and keeping for 15 min before cooling. Peaks were identified by comparison with known standards (Supelco 37 Component FAME Mix). Fatty acids composition (wt %) were calculated by the corrected peak area normalization method. The chromatographic system and the peak integration were managed using Galaxie software (Agilent, Santa Clara, CA).

2.7. Laurdan fluorescence measurements

Laurdan (6‐lauroyl‐2‐dimethylaminonaphthalene) is a fluorescent probe which locates in the bilayer by its lauric acid tail, with the fluorescent moiety localized at the level of the glycerol backbone (Antollini & Barrantes, 1998). It has a very high partition coefficient from aqueous environments to membranes and it is equally distributed between gel and liquid‐crystalline (LC) phases. It is not affected by pH or by the chemical nature of phospholipid polar heads. Its spectral sensitivity to the membrane phase state can be related to its ability to sense polarity and the dynamics of water molecules (dipolar relaxation of water molecules) in the immediate vicinity of the fluorophore (Parasassi, De Stasio, Ravagnan, Rusch, & Gratton, 1991). Laurdan's spectral features are used to calculate the generalized polarization (GP) equation which provides information about the phospholipid phase of the membrane (Bagatolli & Gratton, 1999). Laurdan excitation (Ex) GP spectra were calculated as follows (Parasassi, Loiero, Raimondi, Ravagnan, & Gratton, 1993): Ex GP = I450 − I490/I450 + I490 where I450 and I490 are the intensities at each excitation wavelength, from 320 to 420 nm, obtained using a fixed emission wavelength of 450 and 490 nm, respectively. In the phospholipid gel phase, Laurdan Ex GP spectra are wavelength independent; in the LC phase, Ex GP values decrease with increasing excitation wavelength, with two coexisting phases, the GP spectrum has an opposite trend.

Low Ex GP values indicate a large water molecular mobility around the probe (thus a fluid membrane), while high Ex GP values indicate a rigid membrane with restricted reorientation of water molecules during the fluorescence lifetime. Laurdan steady‐state fluorescence measurements were carried out on a computer‐controlled Perkin Elmer (PerkinElmer Ltd., Buckinghamshire, UK) LS55 spectrofluorimeter. The fluorescence background obtained from cells without Laurdan was always subtracted from the data. Laurdan fluorescence spectra were measured at 37 °C and the temperature was measured in the sample by a digital thermometer. Cells were labeled with Laurdan at a final probe concentration of 1 μM. Each Laurdan spectrum corresponds to the average of determinations performed on five different samples (n = 5). All spectra were normalized by using Perkin Elmer FL WinLab Software.

2.8. RNA isolation and real‐time PCR

Primary myometrial and leiomyoma cells were treated with EPA or DHA (50 μM), and kept negative control (treated with 0.2% ethanol as we used to dissolve EPA and DHA) for 48 h, and lysed using TRIzol reagent (Ambion, Life Technologies), and stored at −80 °C. Total RNA (colorless upper aqueous phase) was separated using chloroform according to the manufacturer's instructions. After that, RNA was purified and concentrated using ReliaPrep™ RNA Cell Miniprep System (Promega Italia, Milan, Italy). The complementary DNA (cDNA) was generated from 1 μg of RNA using high‐capacity cDNA reverse transcriptase (RT) kit (Applied Biosystems, Life Technologies), and newly synthesized cDNA was used for real‐time PCR. Real‐time PCR was performed on StepOnePlus version 2.2.2 (Applied Biosystems, Life Technologies) in 96‐well optical reaction plates with 50 ng cDNA in a final volume of 15 μl, containing 1X TaqMan® fast advanced master mix, with the following TaqMan® gene expression assays (Applied Biosystems, Life Technologies) (Table 1). Controls included RNA subjected to RT‐PCR without reverse transcriptase.

Table 1.

List of primers used in this study

Gene name Gene symbol Gene aliases Reference sequence Assay ID Amplicon length
Collagen, type I, alpha 1 COL1A1 OI4 NM_000088.3 Hs00164004_m1 66
Fibronectin 1 FN1 CIG, ED‐B, FINC, FN, FNZ, GFND, GFND2, LETS, MSF NM_002026.2 NM_054034.2 NM_212474.1 NM_212476.1 NM_212478.1 NM_212482.1 Hs00365052_m1 82
Versican VCAN CSPG2, ERVR, GHAP, PG‐M, WGN, WGN1 NM_001126336.2 NM_001164097.1 NM_001164098.1 NM_004385.4 Hs00171642_m1 72
Inhibin, beta A INHBA EDF, FRP NM_002192.2 Hs00170103_m1 65
Integrin, beta 1 (fibronectin receptor, beta polypeptide, antigen CD29 includes MDF2, MSK12) ITGB1 CD29, FNRB, GPIIA, MDF2, MSK12, RP11‐479G22.2, VLA‐BETA, VLAB NM_002211.3 NM_033668.2 NM_133376.2 Hs00559595_m1 75
Protein tyrosine kinase 2 PTK2 FADK, FAK, FAK1, FRNK, PPP1R71, p125FAK, pp125FAK NM_001199649.1 NM_005607.4 NM_153831.3 Hs01056457_m1 76
A kinase (PRKA) anchor protein 13 AKAP13 AKAP‐13, AKAP‐Lbc, ARHGEF13, BRX, HA‐3, Ht31, LBC, PRKA13, PROTO‐LB, PROTO‐LBC, c‐lbc, p47 NM_001270546.1 NM_006738.5 NM_007200.4 Hs00180747_m1 105
ATP‐binding cassette, sub‐family G (WHITE), member 1 ABCG1 ABC8, WHITE1 NM_004915.3 NM_016818.2 NM_207174.1 NM_207627.1 NM_207628.1 NM_207629.1 Hs00245154_m1 58
ATP‐binding cassette, sub‐family A (ABC1), member 1 ABCA1 ABC‐1, ABC1, CERP, HDLDT1, TGD NM_005502.3 Hs01059118_m1 61
Caveolin 1, caveolae protein, 22 kDa CAV1 BSCL3, CGL3, MSTP085, PPH3, VIP21 NM_001172895.1 NM_001172896.1 NM_001172897.1 NM_001753.4 Hs00971716_m1 66
Sterol regulatory element binding transcription factor 2 SREBF2 CTA‐250D10.14–005, SREBP‐2, SREBP2, bHLHd2 NM_004599.3 NR_103834.1 Hs01081784_m1 91
Cytochrome P450, family 11, subfamily A, polypeptide 1 CYP11A1 CYP11A, CYPXIA1, P450SCC NM_000781.2 NM_001099773.1 Hs00167984_m1 77
Phosphatidylethanolamine binding protein 1 PEBP1 HCNP, HCNPpp, PBP, PEBP, PEBP‐1, RKIP NM_002567.2 Hs00831506_g1 134
Hypoxanthine phosphoribosyltransferase 1 HPRT1 HGPRT, HPRT NM_000194.2 Hs99999909_m1 100
Actin, beta ACT B BRWS1, PS1TP5BP1 NM_001101.3 Hs99999903_m1 171
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2.9. Statistical analyses

Statistical analyses were performed using GraphPad Prism software (version 6.01 for Windows) (GraphPad, San Diego, CA). The data were analyzed using non‐parametric “Kruskal–Wallis” ANOVA, followed by post hoc “Dunn” test for multiple comparisons. Results are expressed as mean  ±  SD. Differences were considered significant when *p < 0.05, **p < 0.01, ***p < 0.001. The data obtained from the GC‐FID analysis of tissues and cells FA were organized and processed by means of Office Excel software (Microsoft). Principal component analysis (PCA) of the results was performed by means the Office Excel add‐in Multibase package (Numerical Dynamics, Japan).

3. RESULTS

3.1. Total fatty acid composition of the leiomyoma and myometrial tissues

The fatty acid composition of the leiomyoma and myometrial tissues (n = 8) is summarized in Table 2. Arachidonic acid was the most abundant fatty acid in both leiomyoma and myometrial tissues, followed by oleic acid, and two saturated species, palmitic and the stearic acids. The fatty acid composition of leiomyoma and myometrial tissues had both a high content of saturated fatty acids (SFA) (> 40%) and a lower content of monounsaturated fatty acids (MUFA) (about 21–22%). The total amount of PUFA was similar in the two tissues, with relative percentages of approximately 35% for each.

Table 2.

Total fatty acid composition (weight % of total fatty acids) of the leiomyoma (FT) and myometrial (MT) tissues, with statistical analysis

Name Carbon number: double bond number Weight % of total fatty acids p
FT MT
Myristic acid 14:0 2.2 ± 0.7 2.0 ± 0.2
Pentadecylic acid 15:0 1.6 ± 0.3 2.0 ± 0.6
Palmitic acid 16:0 17.6 ± 1.5 17.5 ± 1.4
Margaric acid 17:0 0.3 ± 0.1 0.3 ± 0.1
Stearic acid 18:0 16.9 ± 1.4 17.4 ± 0.8
Lignoceric acid 24:0 2.7 ± 0.5 3.1 ± 0.4 *
ΣSFA 41.2 ± 0.8 42.3 ± 1.3 *
Pentadecenoic acid 15:1 0.5 ± 0.2 0.5 ± 0.3
Palmitoleic acid 16:1 n‐7 0.8 ± 0.3 0.7 ± 0.2
Oleic acid 18:1 n‐9 19.0 ± 1.4 18.1 ± 1.8
Vaccenic acid 18:1 n‐7 2.5 ± 0.7 2.2 ± 0.3
Nervonic acid 24:1 0.2 ± 0.2 0.3 ± 0.1
ΣMUFA 22.3 ± 1.1 21.1 ± 1.6
Linoleic acid 18:2 n‐6 cis 8.0 ± 1.4 5.9 ± 0.7 ***
γ‐Linolenic acid 18:3 n‐6 0.2 ± 0.1 0.1 ± 0.0
α‐Linolenic acid 18:3 n‐3 0.1 ± 0.0 0.1 ± 0.0
Stearidonic acid 18:4 n‐3 0.5 ± 0.2 0.5 ± 0.1
Eicosadienoic acid 20:2 n‐6 0.3 ± 0.1 0.3 ± 0.1
Dihomo‐γ‐linolenic acid 20:3 n‐6 3.2 ± 0.4 3.4 ± 0.8
Arachidonic acid 20:4 n‐6 19.9 ± 1.7 22.4 ± 1.5 **
Eicosapentaenoic acid 20:5 n‐3, EPA 0.2 ± 0.1 0.2 ± 0.1
Docosapentaenoic acid 22:5 n‐3, DPA 1.1 ± 0.2 1.0 ± 0.2
Docosahexaenoic acid 22:6 n‐3, DHA 1.2 ± 0.2 1.3 ± 0.4
ΣPUFA 34.7 ± 1.0 35.1 ± 1.5
ΣPUFA ω3 3.0 ± 0.3 3.0 ± 0.5
ΣPUFA ω6 31.6 ± 1.6 32.1 ± 1.3
Lipophilic index 20.6 ± 1.0 20.1 ± 1.5
Other peaks 0.9 ± 0.3 0.7 ± 0.4
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SFA, saturated fatty acid; MUFA, monounsaturated fatty acid; PUFA, polyunsaturated fatty acid. Results represent means ± S.D. (n = 8);. − p > 0.05 (no significant difference); * p < 0.05; ** p < 0.01; *** p < 0.001.

Although the fatty acid composition of leiomyoma and myometrial tissues was highly similar, some statistically significant differences were apparent. For example, the myometrial tissue was significantly richer in arachidonic acid (22.4 ± 1.5%) than the myometrial tissue (19.9 ± 1.7%). Also, the leiomyoma tissue had a higher content of linoleic acid, the endogenous precursor of arachidonic acid, than myometrial tissue (8.0 ± 1.4% vs. 5.9 ± 0.7%, respectively).

The total SFA percentage of myometrial tissue was significantly higher than that of leiomyoma tissue (42.3 ± 1.3% vs. 41.2 ± 0.8%, respectively), but no significant differences were found between the levels of palmitic and stearic acids, the two main saturated species. Concerning the SFA profile, the only significant difference was that lignoceric acid (C24:0) was higher in myometrial than in leiomyoma samples (3.1 ± 0.4% vs. 2.7 ± 0.5%, respectively).

For both myometrial and leiomyoma tissues, the ω6 and ω3 fatty acids comprised approximately 30% and 3% of the total fatty acid content, respectively.

3.2. Effects of DHA and EPA on fatty acid composition of primary myometrial and leiomyoma cells in vitro

The results concerning the fatty acid analysis of primary cells isolated from myometrial tissue and then treated with EPA or DHA in vitro are reported in Table 3. In both treated and untreated myometrial cells (n = 8) the SFA were the main constituents, while the PUFA were present in lower abundance. In the untreated control cells, the principal fatty acid was oleic acid (25.6 ± 2.0%) followed by palmitic acid (19.9 ± 2.3%) and stearic acid (18.0 ± 2.8%). By contrast, palmitic acid was the most abundant fatty acid in cells treated with either EPA (21.8 ± 2.1%) or DHA (21.1 ± 1.4%), followed by stearic acid and oleic acid. Myometrial cells treated with DHA showed a significant reduction of MUFA content compared to the untreated cells and an enhancement of the PUFA fraction, although this increase did not achieve statistical significance. Upon treatment of myometrial cells with EPA, both the MUFA reduction and the PUFA increase were significant respect to control cells.

Table 3.

Total fatty acid composition (weight % of total fatty acids) of cells isolated from myometrial tissue, untreated (Myo_ctrl) and treated in vitro with DHA (Myo_DHA) and with EPA (Myo_EPA), with statistical analysis

Fatty acids Myo_ctrl Myo_DHA Myo_EPA
Name Carbon number: double bond number Weight % of total fatty acids
Myristic acid 14:0 2.4±0.7 2.5±0.5 2.6±0.5
Pentadecylic acid 15:0 1.3±0.2 1.1±0.2 1.1±0.2
Palmitic acid 16:0 19.9±2.3 21.8±2.1 21.1±1.4
Margaric acid 17:0 0.9±0.1 0.8±0.1 0.8±0.1
Stearic acid 18:0 18.0±2.8 19.7±2.4 18.5±2.4
Lignoceric acid 24:0 1.1±0.3b 0.7±0.1a 0.8±0.1a
ΣSFA 43.5±5.3 46.6±4.1 44.9±2.9
Pentadecenoic acid 15:1 1.0±0.8 0.8±0.5 0.8±0.4
Palmitoleic acid 16:1 n‐7 5.7±0.5b 4.3±0.6a 4.2±0.7a
Heptadecenoic acid 17:1 0.6±0.4 0.4±0.3 0.3±0.3
Oleic acid 18:1 n‐9 25.6±2.0b 18.6±3.0a 17.9±2.7a
Vaccenic acid 18:1 n‐7 8.9±2.3b 6.7±1.9ab 6.4±1.6a
Nervonic acid 24:1 0.3±0.1b 0.1±0.0a 0.1±0.0a
ΣMUFA 42.1±1.1b 30.8±4.9a 29.7±4.7a
Linoleic acid 18:2 n‐6 cis 1.0±0.4a 1.6±0.3b 1.6±0.3b
γ‐Linolenic acid 18:3 n‐6 0.0±0.0 0.1±0.0 0.1±0.1
α‐Linolenic acid 18:3 n‐3 0.1±0.1 0.1±0.0 0.1±0.0
Stearidonic acid 18:4 n‐3 0.4±0.1b 0.2±0.0a 0.1±0.0a
Eicosadienoic acid 20:2 n‐6 0.1±0.1 0.1±0.0 0.1±0.0
Dihomo‐γ‐linolenic acid 20:3 n‐6 1.1±0.2 1.2±0.3 1.1±0.2
Arachidonic acid 20:4 n‐6 5.2±1.5b 3.4±0.6a 3.4±0.6a
Eicosapentaenoic acid 20:5 n‐3, EPA 0.3±0.2a 1.2±0.8a 5.1±2.2b
Docosapentaenoic acid 22:5 n‐3, DPA 1.7±0.9a 2.2±1.1a 10.5±3.4b
Docosahexaenoic acid 22:6 n‐3, DHA 2.3±1.2a 11.0±5.6b 1.4±0.8a
ΣPUFA 12.3±4.3a 21.0±8.4ab 23.7±7.0b
ΣPUFA ω3 4.8±2.3a 14.7±7.3b 17.4±6.0b
ΣPUFA ω6 7.5±2.1 6.3±1.1 6.4±1.0
Lipophilic index 30.8±5.0 27.1±7.0 23.2±5.9
Other peaks 2.0±0.2 1.6±0.2 1.7±0.3
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SFA, saturated fatty acid; MUFA, monounsaturated fatty acid; PUFA, polyunsaturated fatty acid. Results represent means ± S.D. (n = 8); differences were considered significant for p < 0.05.

Different superscripts (a,b) denote significant differences (p < 0.05), according to the MSD (minimum significant differences) with respect to the other groups.

∑SFA, total weigth % of the saturated fatty acids; ∑MUFA, total weigth % of monounsaturated fatty acids; ∑PUFA, total weigth % of polyunsaturated fatty acids.

With regard to monounsaturated fatty acids, treatment of myometrial cells with DHA caused a reduction in the content of oleic acid (25.6 ± 2.0% to 18.6 ± 3.0%), palmitoleic acid (5.7 ± 0.5% to 4.3 ± 0.6%), and nervonic acid (0.3 ± 0.1% to 0.1 ± 0.0%) relative to control cells. The content of vaccenic acid (18:1 ω7) also appeared to be reduced upon DHA treatment, but this did not reach statistical significance. Treatment with EPA had a similar effect on the polyunsaturated fatty acid composition of myometrial cells and caused the relative content of oleic acid to decrease from 25.6 ± 2.0% in untreated cells to the 17.9 ± 2.7% in treated cells. Also, palmitoleic, nervonic, and vaccenic acids decreased significantly following treatment with EPA.

The PUFA composition of myometrial cells was strongly affected by omega‐3 treatments. The arachidonic acid content decreased from a value of 5.2 ± 1.5% in the control samples to values of 3.4 ± 0.6% following treatment with either DHA or EPA. By contrast, DHA and EPA both increased linoleic acid content from 1.0 ± 0.4% to 1.6 ± 0.3%. Treatment of myometrial cells with DHA and EPA caused a small but significant decrease in the level of 18:4 ω3. Considering the amounts of EPA, DHA, and DPA in myometrial cells, the two treatments had different effects. In cells treated with DHA, the EPA, and DPA levels were not different from those of control cells. However, the cells undoubtedly assimilated DHA since the relative DHA content increased from 2.3  ±  1.2% to 11.0 ± 5.6%. On the other hand, myometrial cells treated with EPA showed a significant increase in the levels of EPA (5.1 ± 2.2% to 10.5 ± 3.4%) and DPA (0.3 ± 0.2% to 1.7 ± 0.9%), while the DHA level was not affected by EPA in vitro addition.

Leiomyoma cells (n = 8) responded to treatment with EPA or DHA in manner that was similar to myometrial cells (Table 4). As was observed with myometrial cells, the SFA were the main fatty acids in the treated and untreated leiomyoma cells, representing about 45% of the total fatty acids. Neither EPA nor DHA treatment affected the saturated acidic fraction, except for causing a slight decrease in the level of lignoceric acid.

Table 4.

Total fatty acid composition (weight % of total fatty acids) of cells isolated from leiomyoma tissue, untreated (Fib_ctrl) and treated in vitro with DHA (Fib_DHA) and with EPA (Fib_EPA), with statistical analysis

Fib_ctrl Fib_DHA Fib_EPA
Name Carbon number: double bond number Weight % of total fatty acids
Myristic acid 14:0 2.1 ± 0.5 2.3 ± 0.6 2.5 ± 0.7
Pentadecylic acid 15:0 1.7 ± 0.5 1.5 ± 0.3 1.5 ± 0.6
Palmitic acid 16:0 20.3 ± 1.9 20.9 ± 2.8 20.8 ± 2.1
Margaric acid 17:0 0.7 ± 0.1 0.8 ± 0.1 0.8 ± 0.1
Stearic acid 18:0 18.3 ± 3.3 19.9 ± 2.2 18.7 ± 1.6
Lignoceric acid 24:0 1.1 ± 0.3b 0.7 ± 0.2a 0.8 ± 0.2a
ΣSFA 44.2 ± 5.5 46.1 ± 5.2 45.1 ± 3.4
Pentadecenoic acid 15:1 0.9 ± 0.4 1.3 ± 0.7 1.5 ± 0.8
Palmitoleic acid 16:1 n‐7 6.5 ± 1.0b 4.9 ± 0.6a 4.9 ± 0.8a
Heptadecenoic acid 17:1 0.2 ± 0.2 0.4 ± 0.3 0.4 ± 0.3
Oleic acid 18:1 n‐9 24.0 ± 2.3b 19.0 ± 0.6a 18.9 ± 0.4a
Vaccenic acid 18:1 n‐7 9.2 ± 0.6b 6.5 ± 0.5ab 6.4 ± 0.5a
Nervonic acid 24:1 0.2 ± 0.1b 0.1 ± 0.0a 0.1 ± 0.0a
ΣMUFA 41.0 ± 3.5b 32.1 ± 0.4a 32.1 ± 1.3a
Linoleic acid 18:2 n‐6 cis 1.0 ± 0.2a 1.5 ± 0.1b 1.5 ± 0.2b
γ‐Linolenic acid 18:3 n‐6 0.1 ± 0.0 0.0 ± 0.0 0.1 ± 0.0
α‐Linolenic acid 18:3 n‐3 0.1 ± 0.0 0.1 ± 0.0 0.1 ± 0.0
Stearidonic acid 18:4 n‐3 0.3 ± 0.2b 0.1 ± 0.0a 0.1 ± 0.0a
Eicosadienoic acid 20:2 n‐6 0.1 ± 0.0 0.2 ± 0.2 0.1 ± 0.0
Dihomo‐γ‐linolenic acid 20:3 n‐6 1.1 ± 0.3 1.2 ± 0.1 1.1 ± 0.1
Arachidonic acid 20:4 n‐6 6.6 ± 1.9b 4.6 ± 1.8a 4.0 ± 0.8a
Eicosapentaenoic acid 20:5 n‐3, EPA 0.3 ± 0.1a 1.5 ± 0.7a 5.3 ± 1.8b
Docosapentaenoic acid 22:5 n‐3, DPA 1.6 ± 0.9a 1.6 ± 0.9a 7.0 ± 0.9b
Docosahexaenoic acid 22:6 n‐3, DHA 1.8 ± 1.1a 9.4 ± 2.9b 1.3 ± 1.0a
ΣPUFA 12.9 ± 2.6a 20.3 ± 4.9ab 20.7 ± 2.1b
ΣPUFA ω3 4.1 ± 2.2a 12.8 ± 4.2b 13.9 ± 2.0b
ΣPUFA ω6 8.8 ± 1.9 7.5 ± 1.9 6.8 ± 0.8
Lipophilic index 30.5 ± 4.2 27.2 ± 5.7 25.4 ± 3.1
Other peaks 1.8 ± 0.4 1.5 ± 0.4 1.7 ± 0.2
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SFA, saturated fatty acid; MUFA, monounsaturated fatty acid; PUFA, polyunsaturated fatty acid. Results represent means ± S.D. (n = 8); differences were considered significant for p < 0.05.

Different superscripts (a,b) denote significant differences (p < 0.05), according to the MSD (minimum significant differences) with respect to the other groups.

∑SFA, total weigth % of the saturated fatty acids; ∑MUFA, total weigth % of monounsaturated fatty acids; ∑PUFA, total weigth % of polyunsaturated fatty acids.

Treatment of leiomyoma cells with DHA caused a reduction in the total MUFA content from 41.0 ± 3.5% in untreated cells to 32.1 ± 0.4% (p < 0.05) in treated cells. EPA treatment similarly caused the MUFA content to drop to 32.1 ± 1.3% and also caused a significant decrease in the relative content of oleic, palmitoleic, vaccenic, and nervonic acids (Table 4). DHA addition reduced the percentages of the same MUFA species, but did not cause a significant decrease in the level of vaccenic acid.

Treatment with either EPA or DHA increased the level of total PUFA in leiomyoma cells, but this increase was significant only for the cells treated with EPA (p < 0.05). As observed for the myometrial cells, EPA addition caused a significant decrease in the levels of arachidonic acid (6.6 ± 1.9% to 4.0 ± 0.8%) and of C18:4 ω3 (0.3 ± 0.2% to 0.1 ± 0.0%). On the contrary, EPA significantly increased linoleic acid, EPA, and DPA content.

Beyond the significant reduction of the levels of arachidonic and C18:4 ω3, the increase in the content of linoleic acid (p < 0.5) was similar to that caused by the addition of EPA. In contrast to the EPA effect, treatment of leiomyoma cells with DHA caused a significant increase in DHA levels (1.8 ± 1.1% vs. 9.4 ± 2.9%), with no effect on DPA and EPA content. The total PUFA ω3 content of leiomyoma cells was also enhanced from 4.1 ± 2.2% in untreated cells to 12.8 ± 4.2% in DHA treated cells and 13.9 ± 2.0% in EPA treated cells. DHA and EPA treatments had not effect on the PUFA ω6 fraction.

Considering our data (Tables 3 and 4), it is noteworthy that cells isolated from myometrial and leiomyoma tissues had similar responses to treatment with either EPA or DHA. Both treatments provoked a reduction in the MUFA content of the cells and an increase in the PUFA content, and neither affected the fraction of saturated FA. The only differences between the cells treated with EPA and those treated with DHA were found in the ω3 FA composition. The cells treated with DHA were enriched in DHA, with no effects on EPA and DPA. By contrast, treatment with EPA caused an increase in DPA and EPA levels in both myometrial and leiomyoma cells. It is interesting that the concentration of DPA in the cells treated with EPA was mainly increased relative to the level of EPA in the cells.

3.3. Effect of EPA and DHA on the membrane architecture of myometrial and leiomyoma cells

Laurdan Ex GP spectra indicated that the cell membranes of both untreated myometrial and leiomyoma cells (n = 5) were in the liquid‐crystalline phase. In both cell types, EPA and DHA treatment resulted in Laurdan Ex GP values that were higher than those of untreated cells, indicating that the probe was in a more rigid environment with a lower rate of mobility due to a less fluid membrane (Figure 1).

Figure 1.

Figure 1

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(a) Laurdan excitation generalized polarization (Ex GP) spectra in myometrial cells; untreated (—) and treated with EPA (− · −) or DHA (— —). (b) Laurdan excitation generalized polarization (Ex GP) spectra in leiomyomas cells that were untreated (‐‐‐) or treated with EPA (− · · −) or DHA (· · · ·)N = 5

3.4. Effect of EPA and DHA on mRNA expression of extracellular matrix components and activin A in primary myometrial and leiomyoma cells

EPA or DHA (50 µM for 48 hr) treatment had no significant effect on collagen1A1, fibronectin, versican, or activin A mRNA expression in myometrial and leiomyoma cells (n = 3), compared to the untreated control (Figure 2).

Figure 2.

Figure 2

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Effect of EPA and DHA treatment on mRNA expression of extracellular matrix components and activin A in primary myometrial and leiomyoma cells. Data are expressed as mean ± SD (n = 3). NT, No treatment; EPA, Eicosapentaenoic acid; DHA, Docosahexaenoic acid

3.5. Effect of EPA and DHA on mRNA expression of mechanical signaling molecules in primary myometrial and leiomyoma cells

The effects of EPA or DHA treatment on the expression of mechanical signaling molecules, such as integrin β1, FAK, and AKAP13 in myometrial and leiomyoma cells are shown in Figure 3. The real‐time PCR showed that AKAP13 was significantly downregulated by EPA and by DHA about 0.8 fold changed over control, in both myometrial and leiomyoma cells (Figure 3). In addition, in myometrial cells, FAK mRNA expression was decreased by EPA and DHA about 0.8 fold changed over control (Figure 3).

Figure 3.

Figure 3

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Effect of EPA and DHA treatment on mRNA expression of mechanical signaling molecules in primary myometrial and leiomyoma cells. Data are expressed as mean ± SD (n = 6). *p < 0.05; **p < 0.01. NT, No treatment; EPA, Eicosapentaenoic acid; DHA, Docosahexaenoic acid

3.6. Effect of EPA and DHA on mRNA expression of sterol regulatory molecules in primary myometrial and leiomyoma cells

Results of real‐time PCR, performed on primary myometrial and leiomyoma cells treated with EPA or DHA at 50 µM for 48 hr, showed that ABCG1 and ABCA1 were significantly reduced by EPA and by DHA to about one fifth of the untreated control, in both myometrial and leiomyoma cells (Figure 4). In addition, in myometrial cells, SREBF2 was downregulated 0.72 fold change by DHA treatment compared to untreated control (Figure 4).

Figure 4.

Figure 4

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Effect of EPA and DHA treatment on mRNA expression of sterol regulatory molecules in primary myometrial and leiomyoma cells. Data are expressed as mean ± SD (n = 9). *p < 0.05; **p < 0.01; ***p < 0.001. NT, No treatment; EPA, Eicosapentaenoic acid; DHA, Docosahexaenoic acid

3.7. Effect of EPA and DHA on mRNA expression of mitochondrial enzyme CYP11A1 in myometrial and leiomyoma cells

Considering the role of CYP11A1 in the catabolism of steroid hormones, we aimed to see if treatment with EPA and DHA alters the expression of CYP11A1 in myometrial and leiomyoma cells. We found that CYP11A1 mRNA expression was significantly reduced of about a third by EPA and DHA, in myometrial cells but not in leiomyoma cells (Figure 5).

Figure 5.

Figure 5

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Effect of EPA and DHA treatment on expression of CYP11A1 in primary myometrial and leiomyoma cells. Data are expressed as mean ± SD (N = 9). *p < 0.05; **p < 0.01. NT, No treatment; EPA, Eicosapentaenoic acid; DHA, Docosahexaenoic acid

4. DISCUSSION

The lipidomic investigation through GC‐FID analysis revealed the specific fatty acid composition in leiomyoma and healthy myometrial tissue, for example, our results show that arachidonic acid is present at significantly higher levels in myometrial tissue than in leiomyoma tissue, while linoleic acid was higher in leiomyoma tissue. Although not exhaustive, this study represents the first demonstration that uterine leiomyoma tissue differs from the adjacent normal myometrial tissue with regard to its fatty acid profile .

We also tested if it is possible to modulate the fatty acid composition of primary myometrial and leiomyoma cells cultured in vitro. For this purpose, we tested the effect of treating cells with the ω3 fatty acids, EPA, and DHA. The effects of DHA treatment on leiomyoma cells were also similar to its effects on myometrial cells. Both cell types were enriched in DHA following treatment with DHA, and enriched in EPA following treatment with EPA, demonstrating that the cells are able to incorporate the ω3 fatty acids. The treatment with EPA increased also DPA demonstrating the activation of the elongation process of EPA. We found that both EPA and DHA also reduced the amount of the arachidonic acid. Interestingly, the ability of omega‐3 fatty acids to interfere with arachidonic acid metabolism has been reported to be the heart of their anti‐inflammatory effects (De Caterina & Libby, 1996). These results encouraged us to explore the possible effects of ω3 fatty acids on the cell membrane, which plays a critical role in cellular communication and function.

4.1. Our Laurdan Ex

GP results show that myometrial and leiomyoma cell membranes are in the liquid‐crystalline phase when untreated but that most of the Laurdan molecules are located in a less fluid environment in the membranes following treatment with EPA or DHA. This was surprising since we expected that treatment of both cell types with long‐chain PUFA would be compatible with a much more fluid membrane microenvironment. However, our results are consistent with those of Teague, Ross, Harris, Mitchell, and Shaikh, 2013 who showed that DHA, despite its flexible structure, can increase membrane molecular order in primary B cells, EL4 cells and liposomes of varying composition. Furthermore, Kim, Barhoumi, McMurray, and Chapkin, 2014 used Laurdan fluorescence polarization microscopy to show that membrane enrichment with dietary n‐3 PUFA in immunological synapses of CD4+ T cells from DO11.10 T‐cell receptor transgenic mice increases membrane molecular order.

Lipid‐protein interactions are very important in membranes and proteins may affect lipid properties, such as the acyl chain order, the lateral, and transmembrane distribution and the mobility (Nyholm, 2015). Overall, our data suggest that DHA and EPA remodel membrane architecture with possible effects on several cellular properties including cell signaling. We focused on genes that characterize leiomyomas and that could be modulated in their expression by changes in the structure of the plasma membrane. These include molecules of the ECM, which are anchored externally to the plasma membrane and thus involved in mechanical signals. Considering that we studied reproductive tissues, we also checked molecules of mechanisms involved in steroid synthesis and transport of cholesterol.

While we found no changes in gene expression levels of ECM components and activin A, we detected some modification in the expression of molecules involved in mechanical signal transmission such as integrin β1, FAK, AKAP13, although these changes were mainly restricted to myometrial cells.

Mechanical signals are transmitted from the ECM through integrins, which are heterodimeric transmembrane receptors. Most integrins recruit cytoplasmatic focal adhesion kinase (FAK) and activate downstream pathways, including Rho‐dependent signaling (PaszeK et al., 2005). A kinase anchor protein 13 (AKAP13) is a RhoA GTPase‐specific guanine exchange factor (Rho‐GEF) that converts RhoA from its inactive GDP‐bound form to its active GTP‐bound form. The levels of integrin β1 (Chen, Lin, Cheng, & Wing, 2013), AKAP13 (Rogers et al., 2008), and phosphorylated FAK (Chen et al., 2013) were all found to be higher in leiomyoma relative to myometrium.

We found that treatment of cells with ω3 fatty acids reduces the expression of key genes that mediate lipid accumulation, such as ABCG1 and ABCA1. Those proteins are involved in the efflux of cholesterol, the common precursor of steroid hormones and fat‐soluble vitamins (A, D, E, K) (Ikonen, 2008), as well as a modulator of human oxytocin receptor membrane activity, stability and affinity to oxytocin. In this regard, altered myometrial cholesterol content has been associated with abnormal oxytocin‐induced uterine smooth muscle contraction in rodents. In addition, prevention of accumulation of cholesteryl esters in mouse myometrium by nuclear liver X receptor‐β (LXRβ) occurs via regulation of ABCA1 and ABCG1 (Mouzat et al., 2007).

Elevated levels of serum HDL (high‐density lipoprotein)‐cholesterol were found in women with fibroids compared with control patients (Sadlonova et al., 2008), suggesting a role of cholesterol regulatory proteins in the pathogenesis of leiomyoma. Recently Borahay and colleagues reported that simvastatin, which is commonly prescribed to lower high cholesterol levels, is able to inhibit the growth of human uterine fibroid tumors (Borahay et al., 2015). ABCG1 and ABCA1 are member of the evolutionarily conserved family of ATP‐binding cassette cholesterol transporters. ABCG1 plays a pivotal role in cellular cholesterol efflux to HDLs (Wang, Lan, Chen, Matsuura, & Tall, 2004) but not to lipid‐free apolipoprotein A‐I (apoA‐I) (Vaughan & Oram, 2005). In contrast, the efflux of cellular cholesterol and phospholipids to apoA‐I mediated by ABCA1 converts apoA‐I into nascent HDL, which can then act as an efficient acceptor for ABCG1‐mediated cholesterol efflux (Smith et al., 2004). CAV‐1 (Caveolin‐1) is a component of caveolae, a “cave‐like” invagination of the cell membrane. CAV‐1 can interact with ABCG1 and regulate ABCG1‐mediated cholesterol efflux (Gu, Wang, & Zhang, 2014). While we found no effect of ϖ3 fatty acids on CAV‐1, we found that SREBP‐2 (sterol‐regulatory‐element‐binding protein‐2) was downregulated in myometrial cells. SREBP‐2 is best known for its regulatory role in cholesterol uptake and cholesterol synthesis. SREBP‐2 can act as a positive regulator of ABCA1 gene expression (Wong, Quinn, & Brown, 2006). The overexpression of ABCG1 increases the processing of SREBP‐2 to the transcriptionally active protein, thus increases in the expression of SREBP‐2 target genes and cholesterol synthesis (Tarr & Edwards, 2008).

CYP11A1 is a mitochondrial enzyme that catalyzes conversion of cholesterol to pregnenolone. Pregnenolone is the precursor of estrogens, progestogens, mineralocorticoids, glucocorticoids, and androgens, as well as the neuroactive steroids. It is well known that estrogen and progesterone have tremendous effect on uterine leiomyoma growth (Ciarmela et al., 2011; Islam et al., 2013).

Our data showed that treatment with ω3 fatty acids seems to have a greater effect on gene expression levels in myometrial cells, while in leiomyoma cells this treatment seems to reduce the ability to modulate the gene expression. In myometrial cells, omega‐3 fatty acids were also able to reduce CYP11A1, the mitochondrial enzyme that catalyzes the conversion of cholesterol to pregnenolone, the precursor of steroid hormones, suggesting that this cellular type differs in the mechanism of regulation of steroidogenesis and this may contribute to the pathogenesis of uterine leiomyoma.

In conclusion, omega‐3 fatty acids modulate lipid profile, remodel membrane architecture, and down regulate expression of genes involved in mechanical signal and cellular lipid accumulation in myometrial cells.

The present study adds the lipidomic approach in leiomyoma characterization in addition to genomic, epigenetic, and proteomic studies, and highlights new insights on molecular mechanisms underlying the pathogenesis of the disease. This new understanding and the demonstration of the possibility to modulate the lipid content will promote clinical applications in drug discovery and therapy.

CONFLICTS OF INTEREST

The authors have nothing to declare.

Islam MS, Castellucci C, Fiorini R, et al. Omega‐3 fatty acids modulate the lipid profile, membrane architecture, and gene expression of leiomyoma cells. J Cell Physiol. 2018;233: 7143–7156. 10.1002/jcp.26537

Md Soriful Islam, Clara Castellucci, and Rosamaria Fiorini contributed equally to this work.

Contributor Information

Deborah Pacetti, Email: d.pacetti@univpm.it.

Pasquapina Ciarmela, Email: p.ciarmela@univpm.it.

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