Nuclear magnetic resonance spectrometers: For the structure determination of **1** and **2**, a Bruker 800 MHz NMR spectrometer (Bruker Biospin—Billerica, Merating with a 5 mm TCI He Cryoprobe was used. 1D 1H-NMR and magnitude 2 D NMR experiments were run apart from the 1H-13C HMBC of **1**, where an additional phase sensitive experiment was conducted. In the case of compound **3**,1D 1H-NMR and magnitude mode 2 D NMR experiments were run including a selective NOESY experiment with a Bruker 600 MHz NMR spectrometer (Bruker Biospin—Billerica, MCE III HD operating with a N2 Cryoprobe. HSQC experiments were obtained using 2D H-1/X correlation via the double inept transfer phase sensitive using Echo/Antiecho-TPPI gradient selection; with decoupling during acquisition; using trim pulses in inept transfer; with multiplicity editing during selection step, shaped pulses for inversion on f2—channel for matched sweep adiabatic pulses; for HMBC, experiments were obtained with 2D H-1/X correlation via heteronuclear zero and double quantum coherence, phase sensitive and magnitude mode in the case of **1**, and magnitude mode for **2** and **3**, using Echo/Antiecho gradient selection with two-fold low-pass *J*-filter to suppress one-bond correlations with no decoupling during acquisition [44–49].

Liquid chromatography-mass spectrometry: **1** and **2** were analyzed with a Bruker MAXIS II qTOF LC-MS instrument. LC-qTOF utilizes a Phenomenex Kinetex XB-C18 column (2.6 μm; 100 × 2.1 mm) with a mobile phase of 5% acetonitrile + 0.1% formic acid to 100% acetonitrile + 0.1% formic acid with a run time of 11 min. **3** was analyzed with a Thermo Instruments MS system (LTQ-XL Orbitrap Discovery) coupled to a 1290 Infinity Agilent UHPLC system, utilizing a Phenomenex Kinetex XB-C18 column (2.6 μm; 100 × 2.1 mm) with a mobile phase of 5% acetonitrile + 0.1% formic acid to 100% acetonitrile + 0.1% formic acid in 25 min. The analytes were diluted to 0.01 mg/mL concentration with methanol.

Infrared spectra were recorded in ethanol with a Perkin Elmer Spectrum Two FTIR spectrometer Perkin Elmer, MVis Spectroscopic data were recorded in ethanol using a Thermo Scientific Evolution 201 UV–Visible Spectrophotometer Thermo, M. Computational Calculations

A Macromodel module implemented in the Maestro Quantum mechanical software (Maestro Schrödinger LLC, New York, N was used for conformational searches [21]. The calculations were carried out by using an OPLS 2005 force field with ethanol as the solvent. A torsional enhanced sampling with 1000 or 10,000 steps was fixed using an energy window of 4.0 kcal/mol. Molecular geometry optimizations were performed at the DFT theoretical level using the Gaussian 09W package with a HSEH1PBE/cc-pVDZ auto for energy and frequency calculations. After removing redundant conformers and those with imaginary frequencies, theoretical Boltzmann energy population-weighted UV–Vis spectra were calculated by using PBEPBE/6-311++g(3d,2p) with 24 states [22]. The open software SpecDis V.1.71 (Berlin, Germany, 2017, https:/specdis-software.jimdo.com, (accessed on the 7th y 2020 and on the 14 April 2021)) was used to obtain the graphical theoretical UV–Vis curves [23].

#### *3.4. Isolation of Compounds*

Dermacozine N (**1**) was obtained with further purification of the initial silica fractions detailed in Section 3.2 as follows: the first fraction (S1) of the initial FD fractionation with silica chromatography gave dermacozine M (**14**) as reported by Abdel-Mageed et al. [8]. The S2–S3 deep purple colored fractions of the initial silica chromatography as described in Section 3.2 were combined based on TLC chromatography, dried in a N2 drier (57.3 mg), and further partitioned with silica glass column chromatography. A Quickfit CR 20/30 19/26 glass column was used, packed with ~20 g silica (~350× of the mass of the initial mass of the combined S2–S3 silica chromatography FD fraction), and the mobile phase was 9:1 dichloromethane:methanol also in this case. In this experiment, fifteen (SA1–SA15) fractions were collected, each into 7 mL vials with a flow rate of ~0.4 mL/min. The SA9– SA11 pink colored fractions of this experiment were combined based on TLC. Keeping this fraction in a cold room at 4 ◦C, a pink precipitate appeared, which was centrifuged at 10,000 rpm for 15 min. Upon removal of the supernatant and drying the substance in N2 drier, the substance appeared to be a pure compound (1.6 mg) according to the 1H-NMR and the LC-HRESI-MSn measurements (Figures S1 and S6).

Dermacozine O (**2**) was obtained with further purification of the initial silica fractions detailed in Section 3.2 as follows: the S5–S8 red colored silica fractions (28.2 mg) were combined based on TLC chromatography, dried in a N2 drier, and subjected to HPLC purification. High performance liquid chromatography purification of dermacozine O (2) was carried out isocratically with a methanol/H2O 60%/40% + 0.1% trifluoroacetic acid solvent system over 40 min with 1.8 mL/min flowrate, and with Sunfire C18 reversed phase column (5 μm; 110 × 250 mm). Peak detection and UV–Vis trace analysis was carried out at the wavelengths of 254 ± 5 nm, 280 ± 5 nm, 330 ± 5 nm, 350 ± 5 nm, and 530 ± 25 nm. Material collection was automated based on UV–Vis peak appearance versus time. The HPLC fraction with RT = 31.9 min yielded (3.1 mg) the pure compound (see Figures S13, S16 and Theme S3).

Dermacozine P (**3**) was obtained with further purification of the initial silica fractions detailed in Section 3.2 as follows: the initial S4 (7 mg) blue colored silica fraction of the dichloromethane Kupchan fraction (FD) was dried in N2 drier and subsequently repurified with High performance liquid chromatography. The HPLC purification of dermacozine P (**3**) was carried out with a gradient from the initial solvent ratio of 0% methanol and 95% H2O + 5% methanol + 0.05% trifluoro-acetic acid to 100% methanol and 0% H2O over 40 min with 2 mL/min flowrate, and with an ACE C18 HL reversed phase column (5 μm; 110 × 250 mm). Peak detection and the UV–Vis trace analysis was carried out at the wavelengths of 250 ± 5 nm, 280 ± 5 nm, 500 ± 25 nm, 550 ± 25 nm, and 600 ± 25 nm. Material collection was automated based on UV–Vis peak appearance versus time. The HPLC fraction with RT = 15.9 min yielded (3.4 mg) the pure compound (see Figures S24, S26 and Theme S4).

#### *3.5. Cytotoxic Activity of Dermacozines against Human Tumor Cell Lines*

The cytotoxic activity of dermacozines N (**1**) and O (**2**) against five different human cancer cell lines, namely A549 (lung carcinoma), A2058 (metastatic melanoma), MCF7

(breast adenocarcinoma), MIA PaCa-2 (pancreatic carcinoma), and HepG2 (hepatocyte carcinoma) (obtained from ATCC, Manasas, V [50] was studied based on the MTT (3-(4,5 dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay [51]. The compounds were tested as a 10-point dose-response curve ( 1 2 serial dilutions) starting at a concentration of 20 μg/mL in triplicate. IC50 values were determined as previously described [50]. Dermacozine P (**3**) was not tested against cancer cell lines.

#### *3.6. Chemical Characterization of Compounds*

Dermacozine N (**1**): pink amorphous powder, 1.6 mg; LC-HRESI-MSn (qTOF, Bruker), *m/z* measured 386.1247 [M + H]+, 408.1068 [M + Na]+, 772.2448 [2M + H]+, 793.2225 [2M + Na]+, Δ = −0.3 ppm for calculated *m/z* of [M + H]+ 386.1248; DBE = 17; MF = C21H15O3N5. UV λmaxEtOH = 729, 660, 577, 364, 309 nm. IREtOH = 3439, 2988, 1662, 1066, 1039, 544, 521, 471, 463 cm−1. 1H- and 13C-NMR data (DMSO-*d*6), see Table 1.

Dermacozine O (**2**): ink blue amorphous powder, 3.1 mg; LC-HRESI-MSn (qTOF, Bruker) *m/z* 398.1125 [M + H]+, Δ = −2.5 ppm for calculated *m/z* of [M + H]+ 398.1135, DBE = 18; MF = C23H15O4N3. UV λmaxEtOH = 644, 563, 463, 396(sh), 308 nm. IREtOH = 2971, 1989, 1949, 1091, 1048, 882, 524, 501, 481, 453 cm−1. 1H- and 13C-NMR data (DMSO-*d*6), see Table 1.

Dermacozine P (**3**): purplish amorphous powder, 3.4 mg; LC-HRESI-MSn (Orbitrap, Xcalibur) *m/z* 372.0990 [M + H]+, Δ = 3.0 ppm for calculated *m/z* of [M + H]+ 372.0979; DBE = 17; MF = C21H13O4N3. UV λmaxEtOH = 465, 415(sh), 363 nm. IREtOH = 3440, 3178, 1661, 872, 757, 603, 575, 556, 534, 523, 465, 437, 410 cm−1. 1H- and 13C-NMR data (DMSO-*d*6), see Table 1.

**Table 1.** NMR spectroscopic data for dermacozine N (**1**) (800 MHz, DMSO-*d*6), dermacozine O (**2**) (800 MHz, DMSO-*d*6), and dermacozine P (**3**) (600 MHz, *DMSO*-*d*6).†: Calculated with ACD Labs software, Nural Nework Algrithm, DMSO-*d*6 (Chemical shift is not observed).

#### *3.7. Regression Models Used in the Structure Elucidation*

We used a linear regression model in the case of dermacozine N (**1**). The experimental\* 13C-NMR chemical shifts of **1** were correlated to the ACD Labs software calculated ones (ACD/Structure Elucidator, version 2019.2.0, Neural Network Algorithm, solvent used for simulation: DMSO-*d*6) in their theoretically possible structures (**A**–**W**). The simple linear regression model showed clear difference between the possible structures keeping with the results of other modalities in the structure elucidation process. The correlation coefficient obtained was R2 = 0.99 (See Section 2.1).

In the case of dermacozine O (**2**), we compared its experimental 13C-NMR chemical shift values to the ones reported in structures **8**, **9**, and **10** [6] due to their structural similarity. The experimentally not observed 13C-NMR shifts (e.g., C-13 and C-15 in **2**) were excluded from the regression analysis, and since in dermacozine M (**14**) C-3 was calculated with ACD Labs software and contains an additional benzoyl ring at C-3, we excluded this structure from the calculations. A multiple regression test was carried out that showed significant similarity between dermacozine O (**2**) and dermacozine E (**8**) based on the *p-*value and *t*-test (see Section 2.2).

In the case of dermacozines P (**3**), linear regression was done between the ACD Labs software calculated (ACD/Structure Elucidator, version 2019.2.0, Neural Network Algorithm, solvent used for simulation: DMSO-*d*6) and its experimentally observed 13C-NMR chemical shift values. The R2 value obtained was R2 = 0.98 (See Section 2.3).

\*The experimental 13C-NMR values for the statistical calculations were measured in DMSO-d6 in the case of structures **1**, **2**, and **3**.

#### **4. Conclusions**

*Dermacoccus abyssi* MT 1.1T is a piezotolerant, halotolerant Actinomycete from the deepest part of the Earth, the Mariana Trench, Challenger Deep. Since the discovery of the strain in 2006, more than a dozen novel, highly colored dermacozines have been isolated from it. This work provides evidence that the bacterium ability of producing new compounds is still not exhausted yet.

Herein, we report on the isolation of three new dermacozines; dermacozine N (**1**), dermacozine O (**2**), and dermacozine P (**3**) when the strain was seed cultured in 20 g/L containing NaCl medium, followed by the cultivation in ISP2 medium, approximating the salinity of its usual habitat of 34.7‰. The difficulty of structure elucidation arising from the polycyclic nature of dermacozines was overcome by the combined approach of utilizing linear/multiple linear regression of experimental and ACD Labs software calculated 13C-NMR chemical shifts, (TD-DFT) UV–Vis spectral simulation, 1D/2D NMR, LC-HRESI-MSn, biosynthetic route considerations, which step by step offered complementary evidence for our structural assignments.

In dermacozine N (**1**)—whose phenoxazine skeleton is unprecedented in nature to the best of our knowledge—the [*b*]benzopyrazine substitution of the core phenoxazine resulted in UV–Vis absorption maxima in the near infrared (NIR) region. Phenoxazines have been reported to exhibit non-linear optical properties; nevertheless the structure of dermacozine O (**2**) is highly similar to those of dermacozines E (**8**), F (**9**), G (**10**), which were suggested to also exhibit non-linear optic properties in computational chemistry studies. This gives a perspective of investigating **1** and **2** in opto-electronics and second harmonic generation. Given the NIR absorption spectrum of **1**, the reported solvatochromic behavior, and bathochromic shift observed in acidic conditions of related phenoxazine skeletons, **1** would be an outstanding candidate to conduct research on in biosensing chemistry, biomolecular labelling, and *in vivo* metabolic mapping at the cellular level. Compound **1** showed weak activity against melanoma (A2058) and hepatocellular human carcinoma cell lines (HepG2) with IC50 values of 51 and 38 μM, respectively. Dermacozine P (**3**) is the oxidized, 5N-demethylated, and C-6 carboxylated derivative of dermacozine B (**5**). Based on the literature data and cytostatic activities of the previously reported dermacozine derivatives and that synthetic modulation of dermacozine-1-carboxamides increased the compounds' anti-tubulin activity, further anti-tumor studies involving synthetic modulation of the compounds of dermacozine N (**1**), O (**2**), and P (**3**) would be promising.

Fascinating research is being conducted in various fields of chemistry to be able to utilize the novelty of these vividly colored compounds including opto-electronics, computational chemistry, biosensing chemistry, and medicinal chemistry exhibiting exciting results. Whilst the potential use of the colorful dermacozines in chemistry keeps stimulating the interest of researchers, their biological role in the mesmerizing depth of the hadal zone remains unknown.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/md19060325/s1, Figure S1: LC-MS analysis of dermacozine N (**1**) (qTOF, Bruker), Figure S2: MS/MS of dermacozine N (**1**) (qTOF, Bruker), Figure S3A: Proposed MS fragmentation pathway of dermacozine N (**1**) showing loss of a (-ON(C6H4)-) fragment, Figure S3B: Mass error between the calculated (Chemdraw modeled) and experimental (qTOF) *m/z* ratio of dermacozine N (**1**), Figure S4: UV–Vis spectrum of dermacozine N (**1**) in EtOH, Figure S6: Dermacozine N (**1**) 1H-NMR spectrum (800 MHz, DMSO-*d*6), Figure S7: 1H-13C HSQC spectrum of dermacozine N (**1**) (800 MHz, DMSO-*d*6), Figure S8: 1H-13C HMBC (magnitude mode) spectrum of dermacozine N (**1**) (800 MHz, DMSO-*d*6), Figure S9: 1H-13C HMBC spectrum of dermacozine N (**1**) with *J* = 2 Hz (800 MHz, DMSO-*d*6), Figure S10: 1H-1H COSY spectrum of dermacozine N (**1**) (800 MHz, DMSO-*d*6), Figure S11: 1H-1H NOESY spectrum of dermacozine N (**1**) (800 MHz, DMSO-*d*6), Figure S12: 1H-15N HMBC spectrum of dermacozine N (**1**) (800 MHz, DMSO-*d*6), Figure S13: LC-MS analysis of dermacozine O (**2**) (qTOF, Bruker), Figure S14: UV-Vis spectrum of dermacozine O (**2**) in EtOH, Figure S15: IR spectrum of dermacozine O (**2**) in EtOH, Figure S16: 1H-NMR spectrum of dermacozine O (**2**) (800 MHz, DMSO*d*6), Figure S17: 1H-13C HSQC spectrum of dermacozine O (**2**) (800 MHz, DMSO-*d*6), Figure S18: 1H-13C HMBC spectrum of dermacozine O (**2**) (800 MHz, DMSO-*d*6), Figure S19: Long Range 1H-13C HMBC correlations of dermacozine O (**2**) with *J* = 2 Hz (800 MHz, DMSO-*d*6), Figure S20: 1H-1H COSY spectrum of dermacozine O (**2**) (800 MHz, DMSO-*d*6), Figure S21: 1H-15N HMBC spectrum of dermacozine O (**2**) (800 MHz, DMSO-*d*6), Figure S22: UV-Vis spectrum of dermacozine P (**3**) in EtOH, Figure S23: IR spectrum of dermacozine P (**3**) in EtOH, Figure S24: LC-MS analysis of dermacozine P (**3**) (Orbitrap, Xcalibur), Figure S25: MS/MS analysis of dermacozine P (**3**) (Orbitrap, Xcalibur), Figure S26: 1H-NMR spectrum of dermacozine P (**3**) (600 MHz, DMSO-*d*6), Figure S27: 1H-13C HSQC spectrum of dermacozine P (**3**) (600 MHz, DMSO-*d*6), Figure S28: 1H-13C HMBC spectrum of dermacozine P (**3**) (600 MHz, DMSO-*d*6), Figure S29: 1H-1H COSY spectrum of dermacozine P (**3**) (600 MHz, DMSO-*d*6), Figure S30: 1D-NOESY spectrum of dermacozine P (**3**) (600 MHz, DMSO*d*6) from irradiation of the signal at 8.03 ppm, Figures S31-S33: 13C-NMR *δ* calculated values of dermacozine A–P (**1**–**14**) using the ACD Labs software with Neural Network Algorithm, solvent DMSO-*d*6, Figures S34-S38: 13C-NMR *δ* calculated values of possible structures (**A**–**W**) of dermacozine N (**1**), using the ACD Labs software with Neural Network Algorithm, solvent DMSO-*d*6, Table S1: Comparison between calculated vs. experimental 13C-NMR *δ* values of dermacozines A–J (**4**–**13**), Figure S39: Linear regression graphics between ACD Labs (Neural Network Algorithm, DMSO-*d*6) calculated vs. experimental 13C-NMR *δ* values of dermacozine A-H (**4**–**11**), Figure S40: Linear regression graphics between ACD Labs (Neural Network Algorithm, DMSO-*d*6) calculated vs. experimental 13C-NMR *δ* values of dermacozine I & J (**12**–**13**), Table S2: Comparison between the experimental 13C-NMR *δ* values of dermacozine N (**1**) vs. the calculated ones of possible structures **A**, **D**, **E**, **G**, **H**, **I**, **J**, **K**, **N**, **P**, **Q**, **V**, **W** by ACD Labs (Neural Network Algorithm, DMSO-*d*6), Figure S41: Linear regression graphics between 13C-NMR *δ* experimental values of dermacozine N (**1**) vs. the ACD Labs calculated ones (Neural Network Algorithm, DMSO-*d*6) (**4**–**13**), Figure S42: Linear regression graphics between 13C-NMR *δ* experimental values of dermacozine N (**1**) vs. the ACD Labs (Neural Network Algorithm, DMSO-*d*6) calculated ones for possible structures of **N**, **P**, **Q**, **V**, **W**, Table S3: Absolute error and the standard deviation of the error from the mean between the calculated (Structure **W**) and the experimental dermacozine N (**1**) 13C-NMR chemical shifts, Table S4: Experimental values of 13C-NMR *δ* chemical shifts of dermacozines E (**8**), F (**9**), G (**9**), and O (**2**) for multiple regression and *t*-test, Figure S43: Multiple regression analysis between the experimental 13C-NMR *δ* values of dermacozines E (**8**), F (**9**), G (**9**) as independent variables and those of dermacozine O (**2**) as dependent variables, Table S5: Residuals between the observed and predicted values of 13C-NMR *δ* of dermacozines E (**8**), F (**9**), G (**9**) as independent variables and those of dermacozine O (**2**) as dependent variables, Table S6: Comparison between experimental and the ACD Labs calculated 13C-NMR *δ* chemical shift values of dermacozines P (**3**) (Neural Network Algorithm, DMSO-*d*6), Figure S44: Linear regression graphics between the experimental 13C-NMR *δ* chemical shifts of dermacozine P (**3**) and the ACD Labs calculated 13C-NMR *δ* values (Neural Network Algorithm, DMSO-*d*6), Table S7: Absolute error and the standard deviation of the error from the mean between the calculated and the experimental dermacozine P (**3**) 13C-NMR chemical shifts, Figure S45: Evaluation of the cytotoxic activity of dermacozine N (**1**) against human (A) Melanoma (A2058) and (B) Hepatocellular carcinoma (HepG2) cell lines (IC50 graphs), Table S8: Experimental NMR spectroscopic data for dermacozine N (**1**) with HMBC, COSY, and NOESY correlations (800 MHz, DMSO-*d*6), Table S9: Experimental NMR spectroscopic data for dermacozine O (**2**) with

HMBC and COSY correlations (800 MHz, DMSO-*d*6), Table S10: Experimental NMR spectroscopic data for dermacozine P (**3**) with HMBC and COSY correlations (600 MHz, DMSO-*d*6), Figure S46: Workflow of dermacozine N (**1**) structure determination, Figure S47: Workflow of dermacozine O (**2**) structure determination, Figure S48: Workflow of dermacozine P (**3**) structure determination, Theme S1: Dermacozine N (**1**) H-9 and NH2-12 distance (modeled with Chemdraw, Chem 3D), Theme S2: TLC plate of the initial FD fraction following Kupchan liquid–liquid partitioning, showing the colorful bands where the dermacozines were isolated from (left) and the same TLC plate under UV 360 nm light (right), Theme S3: HPLC chromatogram for the isolation of dermacozine O (**2**), Theme S4: HPLC chromatogram for the isolation of dermacozine P (**3**).

**Author Contributions:** Conceptualization, B.J., M.J., D.P.-P., C.J.; methodology, B.J., M.J., D.P.-P., C.J., J.N.T., F.R., B.C., K.K.; software, B.J., M.J., D.P.-P., C.J., J.N.T., F.R., B.C.; validation, B.J., M.J., D.P.-P., C.J., F.R., B.C.; formal analysis, B.J., M.J., D.P.-P., C.J., J.N.T., F.R., B.C., K.K.; investigation, B.J., M.J., D.P.-P., C.J., J.N.T., F.R., B.C., K.K.; resources B.J., M.J., D.P.-P., C.J., J.N.T., F.R.; data curation, B.J., M.J., D.P.-P., C.J., J.N.T., F.R., B.C., K.K.; writing—original draft preparation, B.J., M.J., D.P.-P., C.J.; writing—review and editing, B.J., M.J., D.P.-P., C.J., J.N.T., F.R., B.C., K.K.; visualization, B.J., M.J., D.P.-P., C.J., J.N.T., F.R., B.C.; supervision, M.J., C.J., F.R.; project administration, B.J, M.J., D.P.-P., C.J.; funding acquisition, B.J., M.J., D.P.-P., C.J. All authors have read and agreed to the published version of the manuscript.

**Funding:** Dawrin Pech-Puch received his postdoctoral fellowship from the National Council of Science and Technology (CONACYT) of Mexico.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** The authors wish to express their thanks to Michael Goodfellow, School of Natural and Environmental Sciences, Newcastle University for providing the pure colony of *Dermacoccus abyssi* MT 1.1T, to Wasu Pathom-Aree (Chiang Mai University, Thailand), who isolated the strain furthermore to the *Kaiko* operation team and the crew of M.S. Yokosuka, JAMSTEC, Yokosuka, Japan, for the sediment collection. The authors thank and acknowledge the significant contribution of John W. Still—SEM Specialist, ACEMAC Facility, The University of Aberdeen; Lucinda Wight—Microscope and Histology Specialist, School of Medicine, Medical Sciences and Nutrition, The University of Aberdeen for preparing the strain *Dermacoccus abyssi* MT 1.1T for EM imaging; Russel Gray for NMR measurements—The University of Aberdeen, Department of Chemistry, Marine Biodiscovery Centre, NMR Laboratory; Bella Juraj for NMR measurements—The University of Edinburgh, Joseph Black Building, School of Chemistry, NMR Laboratory. Marcel Jaspars and Bertalan Juhasz acknowledge the help of the University of Edinburgh through the EPSR Grant: EP/R030065/1 for the 800 MHz NMR measurements with the 5 mm TCI He Cryoprobe of compounds **1** and **2**. Carlos Jimenéz and Dawrin Pech-Puch acknowledge the help of CESGA for the computational support. Bertalan Juhasz wishes to express his sincere gratitude to his supervisor, Marcel Jaspars, for his invaluable trust and support.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

## *Article* **A Thermotolerant Marine** *Bacillus amyloliquefaciens* **S185 Producing Iturin A5 for Antifungal Activity against** *Fusarium oxysporum* **f. sp.** *cubense*

**Pratiksha Singh 1,†, Jin Xie 1,†, Yanhua Qi 1, Qijian Qin 1, Cheng Jin 1,2,3, Bin Wang 1,2,\* and Wenxia Fang 1,2,\***

**Abstract:** Fusarium wilt of banana (also known as Panama disease), is a severe fungal disease caused by soil-borne *Fusarium oxysporum* f. sp. *cubense* (*Foc*). In recent years, biocontrol strategies using antifungal microorganisms from various niches and their related bioactive compounds have been used to prevent and control Panama disease. Here, a thermotolerant marine strain S185 was identified as *Bacillus amyloliquefaciens*, displaying strong antifungal activity against *Foc*. The strain S185 possesses multiple plant growth-promoting (PGP) and biocontrol utility properties, such as producing indole acetic acid (IAA) and ammonia, assimilating various carbon sources, tolerating pH of 4 to 9, temperature of 20 to 50 ◦C, and salt stress of 1 to 5%. Inoculation of S185 colonized the banana plants effectively and was mainly located in leaf and root tissues. To further investigate the antifungal components, compounds were extracted, fractionated, and purified. One compound, inhibiting *Foc* with minimum inhibitory concentrations (MICs) of 25 μg/disk, was identified as iturin A5 by high-resolution electrospray ionization mass spectrometry (HR-ESI-MS) and nuclear magnetic resonance (NMR). The isolated iturin, A5, resulted in severe morphological changes during spore germination and hyphae growth of *Foc*. These results specify that *B. amyloliquefaciens* S185 plays a key role in preventing the *Foc* pathogen by producing the antifungal compound iturin A5, and possesses potential as a cost-effective and sustainable biocontrol strain for Panama disease in the future. This is the first report of isolation of the antifungal compound iturin A5 from thermotolerant marine *B. amyloliquefaciens* S185.

**Keywords:** antifungal activity; *Bacillus amyloliquefaciens*; Panama disease; *Fusarium oxysporum* f. sp. *cubense*; bioactive compound; iturin A5

#### **1. Introduction**

Crop wastage is primarily affected by plant diseases, which are mostly caused by soil-borne fungi leading to substantial yield reduction and income losses [1]. Banana (*Musa* spp.) is a major cash crop in the tropics and subtropics and is one of the top ten staple foods in the world [2,3]. The majority of vegetatively propagated bananas are susceptible to many pests and diseases, notably Fusarium wilt of banana (FWB). This disease is caused by the soil-borne fungus *Fusarium oxysporum* f. sp. *cubense* (*Foc*) and is recognized as the most damaging and extensively disseminated disease in banana-producing regions worldwide [4,5]. FWB has spread to the world's leading banana-producing areas since the 1990s [6]. In China, FWB has expanded over much in recent years, placing the banana industry's health and long-term sustainability under risk [7]. As the pathogen of FWB,

Q.; Jin, C.; Wang, B.; Fang, W. A Thermotolerant Marine *Bacillus amyloliquefaciens* S185 Producing Iturin A5 for Antifungal Activity against *Fusarium oxysporum* f. sp. *cubense*. *Mar. Drugs* **2021**, *19*, 516. https://doi.org/10.3390/md19090516

**Citation:** Singh, P.; Xie, J.; Qi, Y.; Qin,

Academic Editors: Daniela Giordano and Bill J. Baker

Received: 17 August 2021 Accepted: 8 September 2021 Published: 11 September 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

*Foc* is resistant to stress and capable of living in soil for up to 30 years [8]. *Foc* infects the xylem through the roots, resulting in widespread necrosis, plant mortality, and causing widespread orchard damage [4,6].

Currently, no suitable cultivar has been selected or bred to reduce the occurrence of FWB [9]. This is a worldwide issue with consequences that include losses in a wide variety of banana production methods and plant mortality in certain cases. Crop rotation, the selection of resistant varieties, and chemical or biological approaches are now the most common strategies for managing FWB [10,11]. However, due to the persistent spread of FWB throughout continents, countries, and regions, there is no effective control system [12]. Chemical fungicides are (relatively) the most effective tool for controlling this soil-borne disease [13]. However, scientists have currently found that long-term usage of fungicides might result in negative consequences such as environmental pollution, plant disease outbreaks, escalating production costs due to chemical overuse, and even human toxicity [14]. So far, there has not been a feasible way to manage the *Foc* pathogen in the field.  Therefore, ecologically and environmentally friendly cultural practices are urgently required [16]. In present agriculture, the biocontrol approach has been being investigated for several years, and many helpful bacteria have been proposed for controlling crop diseases. In addition, biocontrol is the only alternative disease-management strategy that explores plant growth-promoting (PGP) antagonistic bacteria or fungi that are regarded to be safe, ecologically friendly, and cost-effective [11,17], whereas many biocontrol agents are selective to host species, pathogen type, environmental circumstances, soil types, seasons, etc. [18]. Hence, a comprehensive study is necessary to find novel biocontrol agents that can respond to a wide range of environments [19].

The marine ecosystem is composed of marine invertebrates, plants, and their related microbial communities, providing a potential source for new bioactive compounds [20]. The marine microorganisms in the marine ecosystem produce the majority of secondary metabolites which have a wide range of functions, including antiviral, antifungal, immunesuppressive, anti-inflammatory, antitumor, and many other biotechnological and pharmaceutical applications [21,22]. The trend of discovering novel compounds from marine microbes has been increasing during the past decade [23]. Several strains of *Bacillus* spp., which have a suppressive effect against plant diseases caused by soil-borne phytopathogens, have expanded rapidly, and interested readers now have vast quantities of information on biocontrol strategies, as well as their applications and efficacies under a wide range of situations [24]. *Bacillus* spp. are chosen in agricultural systems due to their capacity to produce endospores that can withstand heat and desiccation, in addition to their capacity to be formed into stable dry powders with long shelf lives [25]. Furthermore, many *Bacillus* species have fast growth rates, common inhabitation of plant root microflora, and the ability to produce a large number of secondary metabolites, which are important in antibiosis against a variety of harmful pathogens [26]. Biocontrol methods using *Bacillus* spp. are divided into two categories: direct antagonism and indirect antagonism. Nutrition and geographical location competition, production of secondary metabolites with antibiosis activity, secretion of hydrolases, and the emission of volatile organic molecules are examples of direct antagonism, whereas indirect antagonism involves changing the variety of the soil or plant microbial population to induce plant tolerance and prevent pathogens [10]. *Bacillus* produces important antagonistic substances, such as surfactin [27], iturins [28], and fengycin [29], which are usually generated by non-ribosomal peptide synthetase (NRPS) and inducing plant eliciting system against infections [30].

In this study, we isolated a *B. amyloliquefaciens* strain S185 from offshore of the South Sea, China. This strain exhibits antagonism activity against *Foc*, possesses PGP traits and has other properties suitable for biocontrol utilization. Further analysis showed that iturin A5 is the main bioactive compound contributing to antifungal activity. To the best of our knowledge, this is the first report on the inhibition of *Foc* growth by compound iturin A5 extracted from a marine *B. amyloliquefaciens* strain, which is a potential biocontrol strain for FWB.

#### **2. Results**

#### *2.1. A Thermotolerant Marine Bacterium S185 Possesses Antagonistic Activity against Foc*

From sediment samples collected from offshore of South Sea, China, we isolated a thermotolerant bacterium S185. This strain is capable of growing at 28 ◦C, 37 ◦C, 42 ◦C, and 50 ◦C, respectively (Figure 1A). We then examined the antagonistic activity of S185 against *Foc*, the phytopathogen of FWB. As shown in Figure 1B, strain S185 displayed strong antifungal activity against *Foc* with an inhibition percentage of 78% on a dual culture plate (Table S1).

**Figure 1.** Thermotolerant property of S185 and its antagonistic activity towards *Foc*. (**A**). Strain S185 was cultured at 28 ◦C, 37 ◦C, 42 ◦C, and 50 ◦C, respectively. After 24 h cultivation plates were taken out for photographing. (**B**). Dual culture plate assay for the screening of strain S185 against *Foc*, (**I**). *Fusarium oxysporum* f. sp. *cubense* (*Foc*) control plate, (**II**). Growth inhibition of *Foc* pathogen by S185.

Phylogenetic analysis was conducted to identify the strain S185. After PCR of the 16S *rRNA* product and sequencing analysis, the BlastN program was used to compare the 16S *rRNA* sequence of strain S185 to nucleotide sequences in the NCBI GenBank database. The strain S185 displayed over 97% similarity with the aligned species. A phylogenetic tree was constructed using representative strains from BlastN (Figure 2). The phylogenetic tree indicated that the strain was *Bacillus amyloliquefaciens*.

#### *2.2. S185 Displays PGP Traits and Tolerance to Abiotic Stresses*

Bacterial strains releasing secondary metabolites such as ammonia are helpful in the prevention of fungal infections in plants [31]. The S185 strain indeed showed strong ammonia production in the medium, but was negative for phosphorus (P) solubilization, hydrogen cyanide (HCN), and siderophore production (Table S1). Indole acetic acid (IAA) production is a key property of plant growth-promoting bacteria (PGPB). Strain S185 produces 93.96 ± 2.28 and 57.13 ± 1.32 μg·ml−1 IAA in the presence and absence of tryptophan, respectively (Table S1).

In addition, strain S185 exhibited tolerance to abiotic stresses such as pH of 4–9, temperature of 20–50 ◦C, and salt concentration of 1–5%. The optimum growth of S185 was observed at pH 6, temperature 45 ◦C, and 5% of NaCl (Figure 3). These results confirm that S185 displays PGP traits and tolerance to abiotic stresses.

**Figure 2.** Phylogenetic tree representing the position of strain S185 compared to other strains inside the genus *Bacillus* with *Lactobacillus plantarum* as an outgroup. The tree was created via neighborjoining with a scale bar of 0.01 substitutions per nucleotide.

**Figure 3.** Tolerance of strain *B. amyloliquefaciens* S185 towards abiotic stresses. (**A**). Temperature (20–50 ◦C), (**B**). pH (1–9), (**C**). Salt (1–12%).

#### *2.3. S185 Utilizes Broad Carbon Substrates*

Strains having a broad metabolite tolerance are more appropriate for plant nodulation [32]. Carbon substrates utilization pattern of strain S185 was tested against GNIII Biolog plate. Strain S185 was positive in the assimilation of dextrin, D-turanose, D-salicin, *N*-acetyl-D-glucosamine, *N*-acetyl-β-D-mannosamine, *N*-acetyl-D-galactosamine, *N*-acetyl neuraminic acid, D-fructose, D-galactose, 3-methyl glucose, D-fucose, L-fucose, inosine, Dsorbitol, D-mannitol, D-arabitol, myo-inositol, glycerol, D-glucose-6-PO4, D-fructose-6-PO4, D-aspartic acid, D-serine, glycyl-L-proline, L-alanine, L-arginine, L-aspartic acid, L-glutamic acid, L-histidine, L-serine, lincomycin, niaproof 4, pectin, D-gluconic acid, D-glucuronic acid, glucuronamide, mucic acid, quinic acid, tetrazolium violet, tetrazolium blue, and, L-lactic acid on Biolog GN III plate (Table S2). These findings indicate that the S185 strain is capable of using a wide range of carbon substrates.

#### *2.4. S185 Colonizes in All Tissues of Banana Plants*

The capacity of bacteria to colonize plant tissues is important for disease control and plant growth development. Therefore, colonization of green fluorescent protein (GFP) tagged S185 was assessed in tissues of banana plants by confocal laser scanning microscopy (CLSM). In control plants, without GFP-tagged S185 there was no GFP signal in leaf, stem, and root tissues of plants (Figure 4A,C,E), whereas the GFP-tagged S185 cells were observed as green spots in all plant tissues after 96 h of inoculation (Figure 4B,D,F). The strain S185 exhibited maximum colonization in roots followed by leaves and stems (Figure 4).

**Figure 4.** Confocal laser scanning microscopy (CLSM) showing colonization of *B. amyloliquefaciens* S185 in tissues of banana plants. (**A**,**C**,**E**) are sectioned leaf, stem, and root tissues of banana plants without S185 inoculation, (**B**,**D**,**F**) are sectioned leaf, stem, and root tissues of banana plants after 96 h of inoculation with S185. White arrow marks indicate colonization of strain S185 as small green dots in banana plants.

#### *2.5. Antifungal Activity of Main Active Compound* **1** *Extracted from S185*

Despite the strong antagonistic activity of S185 against *Foc*, we next conducted crude extraction of strain S185 to identify the active components. Guided by activity tracing, butanol extracted components were further fractionated by silica gel column chromatography and gel column chromatography; finally, the main active antifungal compound **1** was isolated from S185 with the yield of 20 mg/L (Figure 5A). Compound **1** displayed the best antifungal activity as shown by the inhibition zone (Figure 5B). At the concentration of 25 μg/disk, compound **1** started to inhibit the growth of *Foc* pathogen with a clear inhibition zone of 7.67 mm (Figure 5B). The diameter of the inhibition zone increased to 9.67 mm, 12.33 mm, 13.33 mm, and 14.33 mm at the 50, 100, 200, and 400 ug/disk of compound **1**, respectively (Table S3; Figure 5B). These results confirm that compound **1** produced by strain S185 has strong antifungal activity against *Foc*.

**Figure 5.** Isolation and antifungal activity of main active compound **1** of strain S185. (**A**) Flow chart showing extraction and separation procedure of compound **1** from strain S185 (**B**). Inhibitory effect of compound **1** on the growth of *Foc* at a different level of concentrations. CK; methanol control.

#### *2.6. Identification of Compound* **1** *as Iturin A5*

The extracted compound **1** from S185 was shown as a white powder and directly identified using high-resolution electrospray ionization mass spectrometry (HR-ESI-MS): m/z [M + H]+, 1057.5671 (Figure 6). Based on this information, the molecular formula of this compound was predicted as C49H76N12O14 (Cacl. for C49H77N12O14+: 1057.5676). 1H nuclear magnetic resonance (NMR) reveals signals of the β-amino acid moiety (800 MHz, DMSO-d6, δ) (Figure S1): 7.11 (ovl, 1H, β-amino acid NH), 3.98 (m, 1H, β-amino acid C3H), 2.32 (d, 2H, β-amino acid C2H2), 1.41 (m, 2H, β-amino acid C4H2), 1.29–1.07 (m, β-amino acid aliphatic CH2), 0.83 (t, *J* = 6.4 Hz, 3H, β-amino acid C15H3). 13C NMR shows signals of (DMSO-d6, δ) (Figure S2): 45.8 (β-amino acid, C3), 42.2 (β-amino acid, C2), 35.0 (β-amino acid, C4), 31.8 (β-amino acid, C13), 29.6 (β-amino acid), 29.5 (β-aminoacid), 29.2 (β-amino acid), 29.1 (β-amino acid), 25.8 (β-amino acid, C5), 22.6 (β-amino acid, C14), 14.4 (β-amino acid, C15). All these data were compared with published NMR data and were consistent with the data for iturin A5 [33]. Thus, compound **1** was identified as iturin A5 (Figure 7, Table S4).

#### *2.7. Iturin A5 Inhibited Spore Germination of Foc*

Despite the antifungal activity from iturin series is known, we assessed the effect of iturin A5 on the growth morphology of *Foc* under an inverted microscope. As shown in Figure 8, compared to the 100% germination rate of *Foc* in the absence of iturin A5, the germination rate of *Foc* decreased to 66% and 37% in the presence of 62.5 μg/mL and 125 μg/mL iturin A5, respectively (Table S5). Moreover, the germ tubes displayed significantly distorted morphology such as ballooned tips and loss of polarity (Figure 8).

**Figure 6.** HR-ESI-MS of antifungal compound **1** isolated from *B. amyloliquefaciens* S185.

**Figure 7.** Structure of compound iturin A5 isolated from *B. amyloliquefaciens* S185.

**Figure 8.** Inhibitory effect of antifungal compound iturin A5 on spore germination of *Fusarium oxysporum* f. sp. *cubense* (*Foc*). We incubated 2 × 105 spores/mL of *Foc* with 0, 62.5 and 125 μg/mL of iturin A5 at 28 ◦C. After 24 h incubation, images were taken under an inverted microscope. Control is DMSO. Scale bar, 40 μm.

#### **3. Discussion**

An essential goal of this study is focusing on the isolation and identification of the most effective biocontrol bacteria and antifungal compounds against *Foc*, the devastating pathogen for FWB. Marine microorganisms have been identified as promising natural sources for the development of biocontrol strains and agents [20]. In marine ecology, unusual circumstances such as extreme temperature and salinity provide a high success rate for the discovery of new and innovative microorganisms as well as their secondary metabolites. More than 20,000 distinct bioactive chemicals have been found in marine fauna and flora to date [34,35]. Previously, different species of *Bacillus* with confirmed biocontrol activity against *Foc* have been reported, including *B. amyloliquefaciens* GKT04 [36,37], W19 [38], NIN-6 [39], and NJN-6 [40], *B. velezensis* HN03 [41], and (iii) *B. subtilis* strains B26 [42], B04, B05 and B10 [43], and N11 [44]. However, an extensive screening of the biocontrol strains from marine bacteria to control FWB pathogen is limited.

*Bacillus* species are useful biocontrol candidates as they possess advantages of forming spores to survive for a long time in extreme environments, producing a wide range of physiologically active secondary metabolites that usually impede the growth of plant pathogens [45], and supporting the development and production of more stable commercial formulations over time from an agro-biotechnological perspective. Combining several *Bacillus* spp. strains is an attractive way to increase biocontrol efficiency under various cropping situations and environmental circumstances, given their adaptability and diverse biocontrol mechanisms. Because of the spore-forming abilities, the *Bacillus* group outperforms non-*Bacillus* species in terms of biocontrol efficiency. Spores can live in a variety of adverse conditions and remain stable during commercial production and maintain tolerance to fungicides [10,46]. *Bacillus*-mediated plant growth promotion is linked to the bacteria's ability to produce phytohormones such as gibberellic acid and IAA, which improve host nutrient absorption and increase plant defense responses to biotic and abiotic stresses [47]. IAA produced by *B. amyloliquefaciens* FZB42 promotes root growth and development of lateral roots, resulting in increased nutrient intake from the rhizosphere [48]. Consistent with previous reports, in this study, we observed IAA production by strain S185 in the presence and absence of tryptophan in the medium. Several bacterial strains produce secondary metabolites such as HCN and ammonia, which are helpful in the management of fungal diseases in a variety of plants [31,49–51]. Similarly, the strain S185 showed strong ammonia production as well as potent antifungal activities against *Foc*, demonstrating the dual role of S185 in controlling FWB.

Being a potential biocontrol strain, one key essential is the colonization capacity of the strain to banana plants. Previously, CLSM of GFP-tagged strains was utilized to illustrate the colonization pattern of *B. megaterium* in rice [52], *B. megaterium* and *B. mycoides* in sugarcane [50]. Similarly, when using this strategy, we observed that the GFP-labelled S185 strain was successfully colonized in all banana plant tissues including leaves, stems, and roots (Figure 4). Previous studies have also suggested that strains with a wider metabolite tolerance are preferable for plant nodulation and growth [32,53]. The BIOLOG metabolic profiling study is a useful technique for identifying microbial diversity using different types of substrates. In this study, carbon utilization pattern of strain S185 was observed on the GNIII Biolog plate and the resulting phenotypes indicated that S185 can use a variety of carbon substrates (Table S2).

Variation in ambient temperatures between crop seasons influenced disease outcomes by affecting the biocontrol agent's ability to kill pathogens [54]. In China, bananas are mostly grown in Guangdong, Guangxi, Hainan, Fujian, Yunnan, and Taiwan. However, diseases and adverse weather conditions continue to pose major obstacles to Chinese banana production. In this study, we found the strain S185 was able to grow between 20–50 ◦C, suggesting that the strain can operate consistently in temporally variable environments, thus making it a suitable biocontrol agent against *Foc* for naturally changing field circumstances. Wei et al. have reported that bacterial strains maintaining their activity in a variety of environments are promising candidates for consistent biocontrol applications [54].

Producing a wide range of secondary metabolites as active components is one of the main properties of *B. amyloliquefaciens*. Therefore, we performed the crude extract from S185 and examined antifungal activity against *Foc*. Not surprisingly, the extract displayed strong antifungal activity, suggesting antimicrobials in the S185 secondary metabolites. Subsequently, a compound **1** purified from the crude extract was identified using HR-ESI-MS (Figure 6). In combination with 13C- and 1H- NMR spectra the compound **1** was identified as iturin A5 (Figure 7).

Iturins are well-known antifungal metabolites generated by *Bacillus* strains [45,55,56], which comprise an amphiphilic peptide ring that is made up of seven chiral amino acids [57]. Iturin A, a member of the iturin family, has a high antibiotic action as well as a broad antifungal spectrum, making it a promising biological control agent for decreasing chemical pesticide use in agriculture [57]. Iturin A has also been found to be a useful drug in human and animal clinical studies because of its broad antifungal range, low toxicity, and low allergic impact [58]. Despite having so many benefits over chemical agents, iturin A has had very few practical uses too far, owing to poor strain productivity and very expensive manufacturing costs. To date, many iturin A producing strains have been identified such as *B. amyloliquefaciens*, *B. licheniformis*, *B. methyltrophicus*, *B. subtilis*, and *B. thuringiensis* [59–63]. However, this is the first report of iturin A5 lipopeptides produced from a thermotolerant marine *B. amyloliquefaciens* S185, which exhibited strong antifungal activity against *Foc*.

Iturin A disrupts the fungal cytoplasmic membrane by interacting mostly with ergosterol molecules, resulting in the formation of transmembrane channels that allow essential ions such as K+ to be released. For example, iturin A has been shown to change the permeability of membranes and the lipid content of *S. cerevisiae* cells [64]. Similarly, Hiradate et al. reported iturin A5 secreted by *B. amyloliquefaciens* RC-2 inhibited the development of mulberry anthracnose caused by the fungus, *Colletotrichum dematium* [33]. We also studied the antifungal activity of purified iturin A5 on the growth of *Foc* pathogen. Different concentrations of iturin A5 produced by S185 was given to test the growth of *Foc*, revealing that MIC of iturin A5 was 25 μg/disk for *Foc*. Earlier, MICs of iturin A for other pathogens were also measured, such as iturin A produced by *B. amyloliquefaciens* PPCB004 inhibited *F. aromaticum* and *Botryosphaeria* sp. with MICs of 1.0 and 1.5 mg/mL [65]. Aberrant conidial and spore germination in fungi was observed when treated by iturins produced by *Bacillus* strains [30,65,66]. Optical and fluorescent microscopy revealed morphological alterations in conidia and significant deformities in *F. graminearum* hyphae treated with iturin A [67]. Wang et al. observed *Phytophthora infestans* mycelia was damaged and had a rough and swollen shape after iturin A treatment [68]. Similarly to previous findings, we also observed the major changes and deformations of *Foc* hyphae after treatment with iturin-A5 produced by S185 (Figure 8).

#### **4. Materials and Methods**

#### *4.1. Collection of Bacterial Strains*

The sediment samples were collected from offshore of South Sea, China. The isolation procedure was described below. Ten grams of sediment from each sample were suspended in 90 mL of autoclaved seawater in a flask and shaken at 100 rpm for 1 h at 25 ◦C. Then, the mixture was serially diluted 10−4 times, and 100 μL of each dilution were distributed on 2216E agar plates. The colony morphology was evaluated by incubating all the plates at 25 ◦C for 2–4 days. Antagonistic activity was tested for all isolated strains against *Foc*. *B. amyloliquefaciens* S185 was chosen for identification. 16s *rRNA* sequence of S185 was submitted to the NCBI GenBank database under accession number MZ333473. All collected strains were stored in 25% glycerol solution at −80 ◦C.

#### *4.2. In Vitro Screening for Antifungal, Plant Growth-Promoting (PGP) Traits, and Abiotic Stress Tolerance*

A dual culture plate method was used to test the antifungal efficacy of strain S185 against *Foc*, following the procedure of Singh et al. [69]. Bacterial strain S185 was spotted 3 cm away from 5 mm diameter of actively growing fungal pathogens spotted in the center point of the yeast extract medium (YAG) (yeast extract-5 g, dextrose-20 g, 1000× trace element-1 mL, agar-18 g, and ultrapure water-1000 mL; 1000×trace element: ZnSO4·7H2O-2.2 g, H3BO3-1.1 g, MnCl2·4H2O-0.5 g, FeSO4·7H2O-0.5 g, CoCl2·5H2O-0.16 g, CuSO4-0.16 g, (NH4)6Mo7O24·4H2O-0.11 g, EDTA-5.0 g, and ultrapure water-100 mL). Plate containing only fungal disk served as the control and was incubated at 28 ◦C for 5–7 days. The percentage of inhibition was measured using the formula; [(R1−R2)/R1] × 100, where R1 is radial growth of the fungal pathogen in control plate and R2 is radial growth of fungal pathogen in the presence of test strain.

PGP traits including P-solubilization, production of IAA, siderophore, ammonia, and HCN by strain S185 were estimated by performing standard protocols of Brick et al. [70], Glickmann and Dessaux [71], Schwyn and Neilands [72], Dey et al. [73] and Lorck [74]. The ability of S185 to solubilize P was tested qualitatively using Pikovskaya medium added with tricalcium phosphate. Strain S185 was inoculated in the plate and observed for the clear hallow zone formation surrounding the isolate after 5–7 days incubation at 30 ◦C. IAA production was evaluated with colorimetric technique in the presence (0.5%) and absence of tryptophan in the medium. Siderophore production of strain S185 was detected on chrome azurol S (CAS) medium and the formation of a halo zone on the medium confirmed siderophore production. For ammonia production, strain S185 was cultured for 72 h in 10% sterile peptone water at 30 ◦C and changes in yellow color by adding 0.5 mL of Nessler's reagent established the ammonia production. HCN production ability of strain S185 was tested on a nutrient broth (NB) medium containing 4.4 g/L glycine to produce hydrocyanic acid. A filter paper soaked with 0.5% picric acid and 2% Na2CO3 was placed on a cover plate, then sealed with Parafilm and incubated at 30 ◦C, and alteration of filter paper color displayed HCN production. All assays were repeated three times with five replications.

Strain S185 was further tested in NB for its capacity to survive under different abiotic stress conditions, such as pH, temperature, and NaCl, and the medium without inoculation was employed as a blank. Temperature tolerance was determined by incubating 0.1 mL fresh bacterial solution of strain S185 in NB medium (5 mL) for 36 h at 20, 25, 30, 35, 40, 45, and 50 ◦C in a shaker incubator at 120 rpm, and O.D. was measured at 600 nm. For pH tolerance, pH of the NB medium was adjusted to 1, 2, 3, 4, 5, 6, 7, 8, and 9 with sterile buffers. Fresh culture of strain S185 was placed into 5 mL of LB broth medium with varied pH levels and incubated at 37 ◦C in an incubator shaker at 120 rpm for 36 h, with growth monitored at 600 nm. In addition, the salt tolerance property of strain S185 was tested in 5 mL of NB medium supplemented with 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, and 12% of NaCl concentration. 0.1 mL bacterial suspension (0.1 mL) was transferred in NB tubes and kept at 37 ◦C at 120 rpm in a shaker incubator and growth was measured after 36 h at 600 nm.

#### *4.3. Phylogenetic Study of Strain S185*

Phylogenetic analysis of strain S185 based on 16S *rRNA* sequences was performed with NCBI GenBank reference sequences. Sequence alignment was performed using the ClustalW and BlastN search algorithms, and closely related sequences were retrieved from NCBI. The Neighbor-Joining approach [75] was used to prepare phylogenetic analysis based on 16S *rRNA* sequence using MEGAX version X [76]. The number of differences method [77] was used to compute the genetic evolutionary distances, and the bootstrap test (1000 replicates) was performed as stated by Felsenstein [78].

#### *4.4. Carbon Utilization Pattern of Strain S185*

Carbon substrates utilization pattern of strain S185 was studied using GENIII BIOLOG(R) Phenotype Micro-ArrayTM plate. Strain S185 was cultivated at 30 ◦C on Luria-Bertani (LB) agar medium, then washed and suspended in an inoculation fluid (IF) to achieve a transmittance of 90–98 percent as per the protocol. The phenotypic fingerprint was formed by incubating a 100 μL of cell suspension in the 96 wells of the GNIII Micro-Plate at 30 ◦C for 48 h. During incubation, the wells' respiration rate increases, allowing the cells to use a variety of carbon sources while growing. Increased respiration causes the tetrazolium dye to be reduced, resulting in purple color. Readings were recorded using an automated BIOLOG(R) Micro-Station Reader following the manufacturer guidelines after incubation.

#### *4.5. Tagging of S185 with GFP-pPROBE-pTetr-TT*

The pPROBE-pTetr -TT plasmid expressing green fluorescent protein (GFP) was provided by the Agriculture College, Guangxi University, Nanning, China. Tissue culture banana plantlets obtained from Guangxi Academy of Agricultural Sciences, Nanning, China were used for this experiment. Freshly grown cultures of S185 and GFP-pPROBEpTetr-TT in LB medium were mixed (1:2) and then incubated for 36–48 h at 32 ◦C with a 160 rpm orbital shaker. Following the incubation, 100 μL of bacterial broth was disseminated on LB agar plate and kept overnight to assess the purity of the tagged strain, as well as validate the tagging using confocal laser scanning microscopy (CLSM). Fluorescent strain S185, when exposed to UV light, was chosen for further investigation.

#### Colonization of S185 in Banana Plantlets

Before bacterial inoculation, banana tissue culture plantlets were rinsed with autoclaved distilled water. Plantlets were placed in an autoclaved glass container comprising 50 mL of MS liquid media (mixture of basal salt and sucrose) at 30 ◦C in a growth chamber. After 3–4 days, plantlets were carefully moved to a new bottle comprising tagged bacterial suspension (∼2.0 × 105/mL cell count). The uninoculated banana plantlets (control) were placed in a growth chamber set at 30 ◦C with a 14 h photoperiod and 60 μmol·m−2·s−1 photon flux density. Plantlets were removed after 72–96 h of growth, rinsed with distilled water, and CLSM was used to check for bacterial colonization. Banana leaf, stem, and root (inoculated and uninoculated) samples were sliced into 1 cm lengths and put on the bridge slide with 10% (*v*/*v*) of glycerol before being detected with CLSM at varying emission rates depending on the intensity of autofluorescence UV light (Leica DMI 6000, Mannheim, Germany) [79].

#### *4.6. Cultivation and Extraction of Antifungal Compounds*

Strain S185 was cultured in 12 L of YAG medium at 25 ◦C for 9 days. The solid fermentation products were cut into small pieces and carefully extracted with ethyl acetate (EtOAc)/methanol (MeOH)/acetic acid (HAc) solution (80:15:5, *v*/*v*/*v*). The extraction was performed three times for the preparation of crude extracts. Crude extracts were dissolved in water and first extracted three times with EtOAc and then extracted three times with n-butanol. We obtained 7.981 g of EtOAc extract and 4.674 g of n-butanol extract after condensation and evaporation with a rotary evaporator. The n-butanol extract residue (4.674 g) was purified by Sephadex LH-20 (Amersham Pharmacia) (MeOH, 2 × 100 cm, 1 mL·min−1) to produce 6 fractions (Fr.1.1-Fr.1.6). Fr.1.2 (592.7 mg) was applied to a silica gel column (Qingdao Marine Chemical Factory) (GF254, 3 × 50 cm, 70 g) using a chloroform: methanol (90:10, 80:20, 70:30) solvent gradient system with ammonia water (0.3%) to generate fractions Fr.1.2.1-Fr.1.2.5. Finally, Fr.1.2.5 (326 mg) was purified by Sephadex LH-20 (MeOH, 2 × 100 cm, 1 mL·min−1) to obtain compound **1** (240.8 mg).

#### 4.6.1. Antifungal Assay of Extracted Compound **1**

Antifungal activity of selected compound **1** was assessed by disk diffusion method against pathogenic fungi *Foc* [80]. The extracted compound was dissolved in methanol and added in 6 mm diameter of paper disks at different levels of concentrations, i.e., 25, 50, 100, 200, and 400 μg/disk, respectively. A disc containing only methanol was used as the control. The dried paper discs were applied onto the surface of YAG plates dispersed

with *Foc* at 28 ◦C for 24 h to 48 h. After incubation, the diameter of the inhibition zone was determined. Each treatment was repeated three times.

#### 4.6.2. Structure Identification of Compound **1**

The Nuclear Magnetic Resonance (NMR) spectra were generated using an Agilent NMR system 800 MHz NMR spectrometer (Agilent Technologies Inc., Colorado Springs, CO, USA) to determine the mass of the antifungal compound **1**. Electrospray ionization mass spectrometry (ESI-MS) and high-resolution electrospray ionization mass spectrometry (HR-ESI-MS) were performed using a Thermo Scientific Orbitrap Elite MS spectrometer (Thermo Fisher Scientific, Waltham, MA, USA) to identify the structure of compound **1** isolated from S185.

#### *4.7. Effect of Purified Iturin A5 on Spore Germination of Foc*

The inhibitory effect of purified iturin A5 on spore germination of *Foc* was performed following the standard EUCAST MIC determination method [81] and observed under an inverted microscope (Leica Microsystems CMS GmbH, Ernst-Leitz-Str. 17–37, D-35578, Wetzlar). *Foc* was cultured on a YAG medium at 28 ◦C for 7 days and then mixed with 10 mL of 0.2% Tween-80 solution. Subsequently, we gently scraped the surface of the fungal colony with a sterile loop, and spores were collected. Finally, the spore concentration was diluted to 2 × 105 CFU/mL. 10 mg iturin A5 was dissolved in 1 mL DMSO to obtain the concentration of 10 μg/μL stock solution. 2 × RPMI1640 medium with 2% glucose was used for spore germination, which was carried out in 96-well plates. Firstly, 300 μL liquid spores were added to the Eppendorf tube and mixed with 270 μL of 2 × RPMI1640 medium with 2% glucose. For control, 30 μL DMSO was added to the Eppendorf tube. For treatment, different concentration of iturin A5 (62.5, 125, 250, and 500 μg/mL) was used. 200 μL of prepared different spore suspensions were added into 96-well plates. After 24 h of incubation at 28 ◦C, spore germination was observed under an inverted microscope.

#### **5. Conclusions**

*Bacillus* strains are potential candidates for controlling FWB because of their wide range of secondary metabolites to inhibit the growth of pathogens as well as colonization and PGP properties. Searching for cost-effective and environmentally friendly *Bacillus* to manage banana pathogenic outbreaks is urgent and practical. In this study, a marine bacterium, *B. amyloliquefaciens* S185, exhibited thermotolerance, IAA, and ammonia production, diverse carbon utilization patterns, and good colonizing capacity in banana plants. In parallel, the S185 strain displayed strong antagonistic activity against *Foc*. The compound iturin A5 from S185 is the main bioactive component for antifungal activity by delaying spore germination and disrupting the polarity establishment of *Foc*. In summary, strain S185 and its iturin A5 lipopeptide is a promising biocontrol strain and agent for FWB. However, field testing of this strain is required to evaluate its efficacy in controlling Foc and promoting banana growth in the future.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/md19090516/s1, Figure S1: The 1H spectra of compound **1** in DMSO-d6. Figure S2: The 13C spectra of compound **1** in DMSO-d6. Table S1: Antifungal and plant growth promoting features of thermotolerant marine *Bacillus amyloliquefaciens* S185. Table S2: List of carbon sources utilized by S185 in GNIII Biolog plate. Table S3: Inhibitory effect of compound **1** extracted from S185 on the growth of *Fusarium oxysporum* f. sp. *cubense*. Table S4: NMR spectroscopic data for compound **1**, iturin A2 and iturin A5 in DMSO-d6. Table S5: Effect of iturin A5 on spore germination rate of *Fusarium oxysporum* f. sp. *cubense*.

**Author Contributions:** Conceptualization, B.W. and W.F.; methodology, P.S. and J.X.; software, J.X.; validation, P.S. and J.X.; formal analysis, P.S. and J.X.; investigation, B.W., C.J. and W.F.; resources, Y.Q. and Q.Q.; data curation, P.S. and J.X.; writing—original draft preparation, P.S. and J.X.; writing—review and editing, P.S., J.X., B.W. and W.F.; visualization, W.F.; supervision, W.F.; project

administration, B.W. and W.F.; funding acquisition, B.W., C.J. and W.F. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by National Natural Science Foundation of China (31960032, 32071279), Guangxi Natural Science Foundation (2020GXNSFDA238008) to W.F., Research Startup Funding of Guangxi Academy of Sciences (2017YJJ026) to B.W., Bagui Scholar Program Fund (2016A24) of Guangxi Zhuang Autonomous Region to C.J.

**Data Availability Statement:** Data sharing does not apply to this article as no new data were created or analyzed in this study.

**Acknowledgments:** We are very grateful to Sugarcane Research Center, Guangxi Academy of Agricultural Sciences, Nanning, China for a colonization study using confocal laser scanning microscopy (CLSM).

**Conflicts of Interest:** All the authors report no conflict of interest relevant to this editorial.

#### **References**

## *Article* **Anti-Food Allergic Compounds from** *Penicillium griseofulvum* **MCCC 3A00225, a Deep-Sea-Derived Fungus**

**Cui-Ping Xing 1, Dan Chen 2, Chun-Lan Xie 1, Qingmei Liu 3, Tian-Hua Zhong 1, Zongze Shao 1, Guangming Liu 3, Lian-Zhong Luo 2,\* and Xian-Wen Yang 1,\***

**Abstract:** Ten new (**1**–**10**) and 26 known (**11**–**36**) compounds were isolated from *Penicillium griseofulvum* MCCC 3A00225, a deep sea-derived fungus. The structures of the new compounds were determined by detailed analysis of the NMR and HRESIMS spectroscopic data. The absolute configurations were established by X-ray crystallography, Marfey's method, and the ICD method. All isolates were tested for in vitro anti-food allergic bioactivities in immunoglobulin (Ig) E-mediated rat basophilic leukemia (RBL)-2H3 cells. Compound **13** significantly decreased the degranulation release with an IC50 value of 60.3 μM, compared to that of 91.6 μM of the positive control, loratadine.

**Keywords:** deep-sea microorganism; fungus; *Penicillium griseofulvum*; anti-food allergy; fungal metabolites; marine natural products

#### **1. Introduction**

For the past decade, the trend to discover new compounds from marine microorganisms continues to rise [1], especially from marine fungi [2,3], which accounted for 68% of the reported new marine natural products in 2019 [4]. Of particular importance is the *Penicillium* species, which are recognized as the richest source for the discovery of biologically important and structurally unique secondary metabolites [5–8].

As our ongoing research for novel and bioactive secondary metabolites from the deep sea-derived microorganisms [8–11], the fungal strain *Penicillium griseofulvum* isolated from the Indian Ocean sediment was selected for a systematic chemical examination. As a result, five carotanes, four naphthalenes, and three viridicatol derivates were obtained [12,13]. A continuous study, however, led to the isolation of 10 new (Figure 1) and 26 known compounds. Herein, we report the isolation, structure elucidation, and biological activity of these compounds.

**Citation:** Xing, C.-P.; Chen, D.; Xie, C.-L.; Liu, Q.; Zhong, T.-H.; Shao, Z.; Liu, G.; Luo, L.-Z.; Yang, X.-W. Anti-Food Allergic Compounds from *Penicillium griseofulvum* MCCC 3A00225, a Deep-Sea-Derived Fungus. *Mar. Drugs* **2021**, *19*, 224. https://doi.org/10.3390/md19040224

Academic Editor: Daniela Giordano

Received: 25 March 2021 Accepted: 12 April 2021 Published: 16 April 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

**Figure 1.** Compounds **1**–**10** from *Penicillium griseofulvum* MCCC 3A00225.

#### **2. Results and Discussion**

Compound **1** was isolated as a white powder. Its molecular formula was established as C17H18N2O4 according to the protonated molecule peak at *m*/*z* 337.1176 [M + Na]+ in its (+)−HRESIMS (High Resolution Electrospray Ionization Mass Spectroscopy) spectrum, requiring ten degrees of unsaturation. The 1H and 13C NMR spectroscopic data (Figures S1 and S2, Table 1) displayed 17 carbons, characteristics of one mono-substituted aromatic unit [*δ*H 7.24 (1H, br t, *J* = 7.4 Hz, H-4), 7.33 (2H, dd, *J* = 7.8, 7.3 Hz, H-3, 5), 7.46 (2H, d, *J* = 7.8 Hz, H-2, 6); *δ*C 127.5 (d × 2, C-2/C-6), 128.3 (d, C-4), 129.1 (d × 2, C-3/C-5), 143.2 (s, C-1)], one ortho-disubstituted benzene moiety [*δ*H 7.16 (1H, td, *J* = 7.6, 1.0 Hz, H-5 ), 7.47 (1H, td, *J* = 7.8, 1.5 Hz, H-4 ), 7.60 (1H, dd, *J* = 7.8, 1.4 Hz, H-6 ), 8.51 (1H, d, *J* = 8.1 Hz, H-3 ); *δ*C 122.4 (d, C-3 ), 124.2 (s, C-1 ), 124.7 (d, C-5 ), 128.8 (d, C-6 ), 132.7 (d, C-4 ), 138.8 (s, C-2 )], one methyl [*δ*H 2.89 (3H, s, 7 -NMe); *δ*C 26.8 (q, 7 -NMe)], two oxygenated methines [*δ*H 4.25 (1H, d, *J* = 2.3 Hz, H-8); 5.16 (1H, d, *J* = 2.0 Hz, H-7) *δ*C 75.6 (d, C-7), 77.8 (d, C-8)], and two carbonyls [*δ*C 171.3 (s, C-7 ), 174.0 (s, C-9)]. In the 1H–1H COSY (Correlation Spectroscopy) spectrum, correlations of H-2 (H-6)/H-3 (H-5)/H-4, H-3 /H-4 /H-5 /H-6 , and H-7 (*δ*H 5.16, d, *J* = 2.0 Hz)/H-8 (*δ*H 4.25, d, *J* = 2.3 Hz) confirmed the two benzene units and deduced another fragment of C-7/C-8. By the HMBC (Heteronuclear Multiple-bond Correlation) correlations of H-7 (*δ*H 5.16) to C-1/C-2/C-6/C-9 and H-6 (*δ*H 7.60)/7 -NMe (*δ*H 2.89) to C-7 , **1** was then assigned a phenylpropionyl moiety and a benzamide groups (Figure 2). However, the limited HMBC correlations hindered the connection of these two fragments. Fortunately, crystals of 1 were obtained. By the single X-ray crystallography (Figure 3), the absolute configuration of **1** was then unambiguously assigned as 2-(2*R*,3*S*dihydroxy-3-phenyl-propionylamino)-*N*-methyl-benzamide, and named penigrisamide.

**Table 1.** 1H (400 MHz) and 13C (100 MHz) NMR spectroscopic data of **1**, **3**, **4**, **8**, and **9** in CD3OD.

**Figure 2.** The key 1H–1H COSY (bold) and HMBC (arrow) correlations of **1**.

**Figure 3.** The X-ray crystallography of **1**.

Compound **2** was afforded as a colorless oil. The molecular formula C19H21N3O4 was deduced from (+)-HRESIMS data (*m*/*z* 378.1418 for [M + Na]+), indicative of eleven degrees of unsaturation. The 1H and 13C NMR spectroscopic data (Figures S7 and S8 from the Supplementary Materials, Table 2) exhibited 19 carbons, including three methyl singlets (one oxygenated), two methylenes, seven methines (five olefinic), and seven nonprotonated carbons (one carbonyl and two ketone groups). These signals were closely similar to those of aurantiomide C (11) [14], except that the terminal amino group in **11** was replaced by the methoxy unit (*δ*C 52.2) in **2**. The assumption was confirmed by the

HMBC correlation of 17-OMe (*δ*H 3.46) to C-17 (*δ*C 173.9). Accordingly, the structure of **2** was determined as 17-deamino-17-methoxylaurantiomide C, and named aurantiomoate C.

**Table 2.** 1H (400 MHz) and 13C (100 MHz) NMR spectroscopic data of **2**, **5**, **6**, **7**, and **10**.

a CD3OD. b DMSO-*d*6.

Compound **3** was obtained as a colorless oil. Its molecular formula was established as C17H27N3O5 on the basis of the protonated molecule peak at *m*/*z* 376.1841 [M + Na]+ in its (+)-HRESIMS spectrum, requiring six degrees of unsaturation. Diagnostic NMR data for 3 suggested the presence of a pyroglutamylleucinmethylester (**20**) [15]. Moreover, the 1H–1H COSY correlation of H2-4 (*δ*H 3.63 m)/H2-5 (*δ*H 2.02 m) and H2-6 (*δ*H 2.18 m, 2.00 m)/H-2 (*δ*H 4.47, dd, *J* = 8.4, 2.8 Hz), with HMBC correlations from H-2 (*δ*H 4.47, dd, *J* = 8.4, 2.8 Hz) to C-4/C-5, and H-6 (*δ*H 2.18 m, 2.00 m) to C-1/C-4, allowed for the presence of another pyroglutamyl moiety. The absolute configuration of **3** was determined by the hydrolysis and derivation using Marfey's reagent, and Nα-(2,4-dinitro-5-fluorophenyl)-l-alaninamide (FDDA) derivatives were compared with the retention times of standard FDDA-amino acids (Figure 4). On the basis of the above evidences, **3** was then assigned as *N*,*N*-pyroglutamylleucinmethylester.

Compound **4** was obtained as a colorless oil. Its molecular formula was established as C12H23NO4 based on the sodium adduct ionic peak at *m*/*z* 268.1526 [M + Na]+ in its positive HRESIMS spectrum, requiring two degrees of unsaturation. Its 1H and 13C NMR spectra were very similar to those of pyroglutamylleucinmethylester (**20**) [15], except for a 2-hydroxy-3-methylbutanoyl unit instead of a pyroglutamyl moiety in **4**. This was confirmed by the 1H–1H COSY correlations of H3-4 (*δ*H 1.00, d, *J* = 7.0 Hz) and H3-5 (*δ*H 0.84, d, *J* = 6.8 Hz) via H-3 (*δ*H 2.07 m) to H-2 (*δ*H 3.86, d, *J* = 3.7 Hz). Via detailed analysis of the HMBC spectroscopic data and using Marfey's method (Figure 5), the absolute configuration of **4** was then assigned as methyl-2*S*-hydroxy-3-methylbutanoyl-L-leucinate.

The molecular formula of **5** was established as C24H32O7 by the ion peak at *m*/*z* 455.2040 [M + Na]+ in its positive HRESIMS. The 1H and 13C NMR spectra exhibited 24 carbons, including three doublets and five singlet methyls, one methoxyl, four methines (two oxygenated and two olefinic), and eleven quaternary carbons (six olefinic and two carbonyl carbons). These signals were closely similar to those of penicyrone A [16] except that the hydroxy (*δ*C 82.6) at the C-9 position in penicyrone A was replaced by the carbonyl (*δ*C 202.8) in **5**. This was confirmed by the HMBC correlations from H-7 (*δ*H 6.38, d, *J* = 1.4 Hz)/H-11(*δ*H 6.28, d, *J* = 1.4 Hz)/H3-19 (1.67, s)/H3-20 (*δ*H 2.02, d, *J* = 1.4 Hz) to *δ*C

202.8. Accordingly, **5** was established to be 9-dehydroxy-9-oxopenicyrone A, and named verrucosidinol A.

**Figure 4.** FDDA derivatives of **3** compared with the retention times of standard FDDA-amino acids.

**Figure 5.** FDDA derivatives of **4** compared with the retention times of standard FDDA-amino acids.

Compound **6** presented a molecular formula of C24H34O8 by positive HRESIMS at *m*/*z* 473.2140 [M + Na]+. Comparison of the 1H and 13C NMR spectra of **6** with those of verrucosidinol (**25**) [17] showed they were very similar except that two olefinic carbons at C-4 and C-5 in **25** were replaced by an epoxy group in **6**. This was evidenced by the HMBC correlations from H3-16 (*δ*H 1.84) to C-1/C-2/C-3, H3-17 (*δ*H 1.61) to C-3/C-4/C-5, and H3-18 (*δ*H 1.28) to C-5/C-6/C-7. Therefore, **6** was established as 4,5-dihydro-4,5 epoxyverrucosidinol, and named verrucosidinol B.

Compound **7** had a molecular formula C17H14O6 as assigned by its positive HRESIMS at *m*/*z* 337.0690 [M + Na]+. Its 1H and 13C NMR spectroscopic data greatly resembled those of helvafuranone [18] except for an additional hydroxy substituent at the C-8 position. By detailed analysis of its 1D and 2D NMR spectroscopic data, **7** was then established as 8-hydroxyhelvafuranone.

Compound **8** gave a molecular formula C10H21NO3 as deduced by the protonated molecule peak at *m*/*z* 202.1504 [M − H]− in its negative HRESIMS spectrum. The 1H NMR spectrum exhibited one methyl doublet at *δ*H 0.96 (3H, d, *J* = 6.2 Hz, H-10), and two methyl singlets at *δ*H 1.12 (3H, s, H-9) and *δ*H 1.16 (3H, s, H-8). The 13C and DEPT spectra revealed the presence of 10 carbons, including three methyls, two methylenes, three methines, and one oxygenated and one carbonyl non-protonated carbon. In the 1H–1H COSY spectrum, correlations were found of H-6 via H-5 to H-4/H-3 and of H-3 to H3-10/H-2. By the HMBC correlations of H2-2 (*δ*H 2.24, dd, *J* = 12.8, 5.2 Hz; 1.98, m) to C-1/C-4/C-10, H-6 (*δ*H 3.21, d, *J* = 9.5 Hz) to C-4/C-7, and H3-8 (*δ*H 1.16, s)/H3-9 (*δ*H 1.12, s) to C-6/C-7, the planar structure of **8** was then established. To determine the absolute configuration of C-6, a dimolybdenum tetraacetate [Mo2(OAc)4]-induced circular dichroism (ICD) experiment was employed. The ICD spectrum exhibited a positive Cotton effect at 310 nm (Figure 6). The sign of the diagnostic band at about 310 nm was correlated to the absolute configuration of the chiral centers in the 1,2-diol moiety. According to the rule proposed by Snatzke, the positive sign suggested a positive torsional angle for the O-C-C-O moiety. It was ascertained that the 6*R*-form could maintain the favored conformation in which the bulkyl moiety and O-C-C-O center stayed away from each other. Based on the above evidence, the structure of **8** was then designated as 6*R*,7-dihydroxy-3,7-dimethyloctanamide.

**Figure 6.** The induced CD spectrum of **8** in DMSO solution of Mo2(OAc)4.

Compound **9** was obtained as a white powder. The molecular formula C11H22O5 was deduced from (+)-HRESIMS data at *m*/*z* 257.1237 ([M + Na]+), indicative of one degree of unsaturation. The 1H NMR spectrum showed a methyl at *δ*H 1.14 (d, *J* = 6.2 Hz, H-10) and a methoxyl at *δ*H 3.65 (s, H-11). The 13C NMR and DEPT (Distortionless Enhancement by Polarization Transfer) data displayed 11 carbons, including one methyl, one methoxyl, five methylenes, three methines, and one carbonyl. In the 1H–1H COSY spectrum, two isolated spin systems were observed as H2-2 (*δ*H 2.49, 2.42)/H-3 (*δ*H 3.79)/H2-4 (*δ*H 1.22)/H2-5 (*δ*H 1.82) and H2-6 (*δ*H 1.21)/H-7 (*δ*H 3.54)/H2-8 (*δ*H 1.61, 1.48)/H-9 (*δ*H 3.94)/H3-10 (*δ*H 1.14). These two fragments could be connected by the HMBC correlations of H2-2 (*δ*H 2.49, 2.42) and H-11(*δ*H 3.65) to C-1 (*δ*C 173.7). Therefore, **9** was established as methyl-3,7,9 trihydroxydecanate.

Compound **10** was obtained as a colorless oil. Its molecular formula was established as C10H18O4 on the basis of the protonated molecule peak at *m*/*z* 225.1109 [M + Na]+ in its positive HRESIMS spectrum, requiring two degrees of unsaturation. The 13C NMR spectrum in association with the DEPT spectrum indicated 10 carbon signals ascribed to one methyl doublet (*δ*c 23.9, C-10), five *sp3* methylenes (*δ*c 42.8, C-2; 32.2, C-4; 24.6, C-5; 32.8, C-6; 46.5, C-8), three *sp3* methines (*δ*c 75.8, C-3; 76.2, C-7; 65.3, C-9), and one carbonyl (*δ*c 175.9, C-1). In the 1H–1H COSY spectrum, a long chain of C-2/C-3/C-4/C-5/C-6/C-7/C-8/C-9/C-10 could be deduced by correlations of H2-2 (*δ*H 2.41)/H-3 (*δ*H 3.76)/H2-4 (*δ*H 1.64, 1.21)/H2-5 (*δ*H 1.84, 1.59)/H2-6 (*δ*H 1.52, 1.21)/H-7 (*δ*H 3.57)/H2-8 (*δ*H 1.48)/H-9 (*δ*H 3.93)/H3-10 (*δ*H 1.13). In the HMBC spectrum, H-3 (*δ*H 3.76) was correlated to C-7 and C-1, which constructed a hexacyclic ring via an ether bond between C-1 and C-7. Accordingly, **10** was established as 9-hydroxy-3,7-epoxydecanoic acid.

By comparison of the NMR and MS data with those published in the literatures, 26 known compounds were determined to be aurantiomide C (**11**) [14], cyclopenin (**12**) [19], (−)-cyclopenol (**13**) [20], (3*S*)-1,4-benzodiazepine-2,5-diones (**14**) [21], 3-benzylidene-3,4 dihydro-4-methyl-lH-l,4-benzodiazepine-2,5-dione (**15**) [22], 3-methyl-3,4-dihydroquinazo line-4-one (**16**) [23], 1,2-dihydro-2,3-dimethyl4(3H)quinazolinone (**17**) [24], *N*,*N'*-1,2-pheny lenebis-acetamide (**18**) [25], aconicarpyrazine B (**19**) [26], pyroglutamylleucinmethylester (**20**) [15], cyclo-(L-Trp-L-Phe) (**21**) [27], fructigenine A (**22**) [28], fructigenine B (**23**) [28], brevicompanine B (**24**) [29], verrucosidinol (**25**) [17], (*S*)-penipratynolene (**26**) [30], (*S*)- 4-(2-hydroxybutynoxy)benzoic acid (**27**) [31], (*S*)-4-(2-hydroxybutoxy)benzoic acid (**28**) (CAS:1357392-03-0), (*S*)-2,4-dihydroxy-1-butyl(4- hydroxy)benzoate (**29**) [32], methyl *p*hydroxybenzeneacetate (**30**) [33], 2-hydroxy phenyl acetic acid (**31**) [34], methyl homogentisate (**32**) [35], 5-hydroxymethyl-furaldehyde (**33**) [36], leptosphaerone A (**34**) [37], 3-methyl-2-penten-5-olide (**35**) [38], and (*R*)-mevalonolactone (**36**) [39].

All isolated compounds (**1**−**36**) were evaluated for their antifood allergic activities in RBL-2H3 cells. Compound **13** showed potent degranulation-inhibitory activity with an IC50 value of 60.3 μM, which was stronger than the commercially available antifood allergy medicine, loratadine (IC50 = 91.6 μM), while **14** and **29** showed weak effects with IC50 values of 167.0 and 134.0 μM, respectively (Table 3).

**Table 3.** Inhibition effects of compounds **1**–**36** on RBL-2H3 cell degranulation (*n* = 3).

a Other compounds, including **1**–**12**, **15**–**28**, and **30**–**36**. b Loratadine was a commercially available anti-food allergic medicine.

#### **3. Materials and Methods**

#### *3.1. General Experimental Procedures and Fungal Fermentation*

*Penicillium griseofulvum*, isolated from a sediment sample of the Indian Ocean at a depth of 1420 m, was deposited at the Marine Culture Collection of China (MCCC) with the accession number MCCC 3A00225. It was cultivated on corn medium in 100 × 1 L Erlenmeyer flasks for 62 days. The detailed general experimental procedures, fungal fermentation, and extraction were reported previously [12].

#### *3.2. Isolation and Purification*

The defatted extract (55.4 g) was separated by column chromatography (CC) over silica gel (500 g) using a CH2Cl2-MeOH gradient (0→100%, 49 mm × 460 mm) to give six fractions (Fr.1−Fr.6). Fr.2 (1.9 g) was subjected to ODS (octadecylsilyl) (H2O-MeOH, 5→100%, 15 × 460 mm, 0.5 L for each fraction) to attain five subfractions (sfrs) (sfrs.2.1– sfrs.2.5). Sfr.2.3 (155.0 mg) was purified by column chromatography on Sephadex LH-20 (100 g) (MeOH, 2.0 × 120 cm, 300 mL) to afford **26** (12.7 mg). Fr.3 (2.1 g) was subjected to column chromatography (CC) on ODS (70 g) (H2O-MeOH, 5→100%, 15 × 460 mm, 0.5 L for each fraction) to attain eleven subfractions (sfrs) (sfrs.3.1–sfrs.3.11). Sfr.3.3 (111.6 mg) was subjected to CC over Sephadex LH-20 (70 g) (MeOH, 2.0 × 120 cm, 300 mL) and silica gel (PE-EtOAc, 2:1, 17 × 305 mm) to yield **35** (25.7 mg). Sfr.3.5 (131.2 mg) was chromatographed on Sephadex LH-20 (100 g) (MeOH, 2.0 cm × 180 cm, 500 mL) resulting in two sub-subfractions (ssfrs) (ssfrs.3.5.1− ssfrs.3.5.2). Ssfr.3.5.1 (3.8 mg) was further purified by HPLC using gradient MeOH-H2O (20→70%, 10 × 250 mm, 4 mL/min) to provide **17** (2.4 mg). Ssfr.3.5.2 (41.0 mg) was purified using preparative TLC (CH2Cl2-Me2CO, 20:1) to give **16** (16.6 mg). Compound **15** (30.7 mg) was isolated from Sfr.3.6 (66.9 mg) by CC over Sephadex LH-20 (70 g) (MeOH, 2.0 × 120 cm, 300 mL). Sfr.3.8 (52.4 mg) was chromatographed on a Sephadex LH-20 (70 g) (MeOH, 2.0 × 120 cm, 300 mL) to give two sub-subfractions (ssfrs) (ssfrs.3.8.1− ssfrs.3.8.2), ssfrs.3.8.1 and ssfrs.3.8.2 were purified by preparative TLC on silica gel (CH2Cl2-MeOH, 20:1) to provide **24** (4.9 mg) and **9** (1.7 mg), respectively. Fr.4 (4.9 g) was subjected to ODS (130 g) (H2O-MeOH, 10→100%, 26 × 310 mm, 1.5 L for each fraction) to obtain twelve subfractions (sfrs) (sfrs.4.1− sfrs.4.12). Compound **12** (216.4 mg) was isolated from sfr.4.1 (304.0 mg) by CC over Sephadex LH-20 (100 g) (MeOH, 2.0 × 180 cm, 500 mL). Sfr.4.2 (906.0 mg) was chromatographed on a Sephadex LH-20 (225 g) column (MeOH, 3.5 × 180 cm, 800 mL) and silica gel (PE-EtOAc, 2:1, 46 × 457 mm) to yield **36** (152.9 mg). Sfr.4.3 (644.0 mg) was fractionated by CC over Sephadex LH-20 (225 g) (MeOH, 3.5 × 180 cm, 800 mL) to attain three sub-subfractions (ssfrs) (ssfrs.4.3.1– ssfrs.4.3.3), ssfr.4.3.3 (78.2 mg) was purified by Sephadex LH-20 (70 g) (MeOH, 2.0 × 120 cm, 200 mL), followed by preparative TLC (CH2Cl2- Me2CO, 10:1) to provide **33** (10.0 mg) and **34** (10.5 mg). Sfr.4.4 (270.9 mg) was subjected to CC over Sephadex LH-20 (100 g) (MeOH, 2.0 × 180 cm, 500 mL), further purified using preparative TLC (PE-EtOAc, 1:2) to obtain **14** (48.6 mg). Sfr.4.5 (33.3 mg) was purified by Sephadex LH-20 (70 g) (MeOH, 2.0 cm × 120 cm, 300 mL) to yield **18** (8.6 mg). Sfr.4.6 (270.9 mg) and sfr.4.7 (342.5 mg) were subjected to CC over Sephadex LH-20 (225 g) (MeOH, 3.5 × 180 cm, 800 mL) to attain **32** (4.0 mg) and **31** (2.9 mg), respectively. Sfr.4.9 and sfr.4.10 (376.6 mg) were fractionated by CC on Sephadex LH-20 (225 g) (MeOH, 3.5 × 180 cm, 800 mL) to obtain four sub-subfractions (ssfrs) (ssfrs.4.10.1–ssfrs.4.10.4). Ssfr.4.10.1 (191.0 mg) was subjected to Sephadex LH-20 (100 g) (MeOH, 2.0 × 180 cm, 500 mL) to attain **22** (117.2 mg), while **28** (3.5 mg) was isolated from ssfr.4.10.3 (10.3 mg) by preparative TLC (CH2Cl2-MeOH, 5:1). Sfr.4.12 (239.5 mg) was chromatographed on Sephadex LH-20 (100 g) (MeOH, 2.0 cm × 180 cm, 500 mL), further purified using preparative TLC (PE-EtOAc, 2:1) to yield **23** (46.2 mg). Fr.5 (40.0 g) separated by column chromatography (CC) over ODS (650 g) (H2O-MeOH, 5→80%, 49 × 460 mm, 3 L for each fraction) to obtain fifteen subfractions (sfrs.5.1–sfrs.5.15). Sfr.5.2 (1.7 g) was separated by CC over Sephadex LH-20 (225 g) (CH2Cl2-MeOH, 1:1, 3.5 × 180 cm, 1000 mL) to give three sub-subfractions (ssfrs) (ssfrs.5.2.1− ssfrs.5.2.3), ssfr.5.2.2 (126.0 mg) was subjected to Sephadex LH-20 (100 g) (MeOH, 2.0 × 180 cm, 500 mL), followed by preparative TLC (CH2Cl2-MeOH, 20:1) to provide **19** (2.9 mg). Ssfr.5.2.3 (103.0 mg) was purified by preparative TLC (CH2Cl2-MeOH, 10:1) to attain **29** (8.6 mg). Sfr.5.3 (625.0 mg) was subjected to CC over Sephadex LH-20 (225 g) (MeOH, 3.5 × 180 cm, 800 mL) to furnish five sub-subfractions (ssfrs) (ssfrs.5.3.1–ssfrs.5.3.5), ssfr.5.3.1(228.0 mg) was separated by silica gel (CH2Cl2-MeOH 50:1→10:1, 46 mm × 305 mm), then subjected to HPLC (MeOH-H2O, 55→65%, 10 × 250 mm, 5 mL/min) to yield **20** (22.8 mg). Compounds **27** (9.3 mg) and **30** (5.1 mg) were isolated from ssfr.5.3.3 (54.0 mg) and ssfr.5.3.5 (29.9 mg) by preparative TLC (CH2Cl2-MeOH, 20:1), respectively, while **10** (3.7 mg) was isolated from ssfr.5.3.4

(29.5 mg) by preparative TLC (EtOAc -MeOH, 50:1), and further purified by preparative TLC (CH2Cl2-MeOH, 20:1). Sfr.5.4 (3.3 g) was fractionated by CC over Sephadex LH-20 (225 g) (3.5 × 180 cm, CH2Cl2-MeOH 1:1, 1200 mL) to attain five sub-subfractions (ssfrs) (ssfrs.5.4.1−ssfrs.5.4.5), ssfr.5.4.2 (73.0 mg) was purified by preparative TLC (CH2Cl2- MeOH, 20:1) to provide **3** (11.0 mg). Ssfr.5.4.3 (1.6 g) was subjected to CC over Sephadex LH-20 (225 g) (3.5 × 180 cm, MeOH, 1200 mL) and preparative TLC (CH2Cl2-MeOH, 20:1) to yield **11** (29.4 mg). Sfr.5.5 (484.0 mg) was subjected to HPLC (MeOH-H2O, 20→40%, 10 × 250 mm, 5 mL/min), followed by preparative TLC on silica gel (CH2Cl2-MeOH, 10:1) to attain **7** (4.5 mg), **13** (34.1 mg), and **8** (4.2 mg). Sfr.5.7 (180 mg) was chromatographed on a Sephadex LH-20 (100 g) (MeOH, 2.0 × 180 cm, 500 mL), and then subjected to preparative TLC (CH2Cl2-MeOH, 20:1) to obtain **1** (1.5 mg). Sfr.5.11 (753.0 mg) was purified by CC over repeated Sephadex LH-20 (225 g) (MeOH, 3.5 × 180 cm, 800 mL) to obtain four sub-subfractions (ssfrs) (ssfrs.5.11.1- ssfrs.5.11.4), **21** (30.9 mg) was isolated from ssfr.5.11.2 (127.5 mg) by preparative TLC on silica gel using CH2Cl2-MeOH (10:1), while **5** (6.1 mg) was isolated from ssfr.5.11.3 (235.7 mg) by preparative TLC on silica gel (PE-EtOAc, 1:1). Sfr.5.12 (3.5 g) was separated by CC over SephadexLH-20 (CH2Cl2-MeOH, 1:1, 3.5 × 180 cm, 1200 mL) to attain three sub-subfractions (ssfrs) (ssfrs.5.12.1–ssfrs.5.12.3). Ssfr.5.12.1 (489.0 mg) was purified by Sephadex LH-20 (225 g) (MeOH, 3.5 × 180 cm, 800 mL) and silica gel (PE-EtOAc, 5:1→1:1, 46 × 305 mm), finally, by preparative TLC (CH2Cl2-MeOH, 10:1) to provide **6** (6.9 mg) and **25** (22.9 mg). Ssfr.5.12.2 (1.6 g) was purified by CC over repeated Sephadex LH-20 (225 g) (MeOH, 3.5 × 180 cm, 1000 mL) and preparative TLC (CH2Cl2-MeOH, 20:1) to yield **4** (3.1 mg) and **2** (24.6 mg).

Penigrisamide (**1**): Colorless needles; [α] 25 *D* +34.5 (c 0.20, MeOH); UV (MeOH) λmax (log ε) 212 (3.03), 252 (2.77) nm; ECD (ACN) Δ*ε*195 +3.67, Δ*ε*203 +1.78, Δ*ε*203 +1.78, Δ*ε*213 +4.40, Δ*ε*225 −0.62, Δ*ε*250 +1.98; 1H and 13C NMR data, see Table 1; (+)-HRESIMS *m*/*z* 337.1176 [M + Na]+ (calculated for C17H18N2O4Na, 337.1164).

*Aurantiomoate C* (**2**): Colorless oil; [α] 25 *D* −20.8 (c 1.20, MeOH), [α] 25 *D* +19.4 (c 1.20, CHCl3); UV (MeOH) λmax (log ε) 211 (4.40), 305 (3.94) nm; ECD (ACN) Δ*ε*191 −20.6, Δ*ε*228 +10.7, Δ*ε*249 −7.66, Δ*ε*272 −1.60, Δ*ε*294 −2.81, Δ*ε*330 +2.17; 1H and 13C NMR data, see Table 2; (+)-HRESIMS *m*/*z* 378.1418 [M + Na]+ (calculated for C19H21N3O4Na, 378.1430).

5-Deoxypyroglutamyl-pyroglutamylleucinmethylester (**3**): colorless oil; [α] 25 *D* −85.6 (c 0.27, MeOH); UV (MeOH) λmax (log ε) 205 (3.77) nm; ECD (ACN) Δ*ε*217 +1.96, Δ*ε*235 −0.39, Δ*ε*249 +0.16; 1H and 13C NMR data, see Table 1; (+)-HRESIMS *m*/*z* 376.1841 [M + Na]+ (calculated for C17H27N3O5Na, 376.1848).

Methyl-2-hydroxy-3-methylbutanoyl-L-leucinate (**4**): colorless oil; [α] 25 *D* −42.9 (c 0.27, MeOH); UV (MeOH) λmax (log ε) 203 (3.31) nm; ECD (ACN) Δ*ε*210 +0.98, Δ*ε*234 −0.11; 1H and 13C NMR data, see Table 1; (+)-HRESIMS *m*/*z* 268.1526 [M + Na]+ (calculated for C12H23NO4Na, 268.1525).

Verrucosidinol A (**5**): Colorless oil; [α] 20 *D* +86.8 (c 0.22, MeOH), [α] 25 *D* +82.7 (c 0.22, MeOH); UV (MeOH) λmax (log ε) 205 (4.13), 231 (4.00), 298 (3.67) nm; ECD (ACN) Δ*ε*187 +1.57, Δ*ε*205 −7.27, Δ*ε*296 +7.91; 1H and 13C NMR data, see Table 2; (+)-HRESIMS *m*/*z* 455.2040 [M + Na]+ (calculated for C24H32O7Na, 455.2046).

Verrucosidinol B (**6**): Colorless oil; [α] 20 *D* + 32.3 (c 0.35, MeOH), [α] 25 *D* +34.6 (c 0.35, MeOH); UV (MeOH) λmax (log ε) 240 (3.98) nm; ECD (ACN) Δ*ε*195 +0.66, Δ*ε*214 −0.89, Δ*ε*254 +4.30; 1H and 13C NMR data, see Table 2; (+)-HRESIMS *m*/*z* 473.2140 [M + Na]+ (calculated for C24H34O8Na, 473.2151).

8-Hydroxyhelvafuranone (**7**)*:* Colorless oil; [α] 25 *D* − 16.7 (c 0.03, MeOH); UV (MeOH) λmax (log ε) 204 (4.28) nm; ECD (MeOH) Δ*ε*193 +2.23; 1H and 13C NMR data, see Table 2; (+)-HRESIMS *m*/*z* 337.0690 [M + Na]+ (calculated for C17H14O6Na, 337.0688).

6,7-Dihydroxy-3,7-dimethyloctanamide (**8**): Colorless oil; [α] 25 *D* −7.3 (c 0.15, MeOH); UV (MeOH) λmax (log ε) 203 (3.09) nm; ECD (MeOH) Δ*ε*225 +0.02; 1H and 13C NMR data, see Table 1; (−)-HRESIMS *m*/*z* 202.1504 [M − H]− (calculated for C10H20NO3, 202.1443).

Methyl-3,7,9-trihydroxydecanate (**9**): White powder; [α] 20 *D* −6.8 (c 0.19, MeOH), [α] 20 *D* −8.9 (c 0.19, CHCl3); UV (MeOH) λmax (log ε) 205 (2.21) nm; ECD (MeOH) Δ*ε*210 +0.11; 1H and 13C NMR data, see Table 1; (+)-HRESIMS*m*/*z* 257.1237 [M + Na]+.

9-Hydroxy-3,7-epoxydecanoic acid (**10**): Colorless oil; [α] 25 *D* +15.7 (c 0.21, MeOH); UV (MeOH) λmax (log ε) 205 (3.10) nm; ECD (MeOH) Δ*ε*211 +0.18; 1H and 13C NMR data, see Table 2; (+)-HRESIMS *m*/*z* 225.1109[M + Na]+ (calculated for C10H18O4Na, 225.1103).

#### *3.3. X-ray Crystallography of 1*

Compound **1** was obtained as colorless needles from MeOH. Its crystallographic data were measured by an Xcalibur and Gemini single-crystal diffractometer with Cu Kα radiation (*λ* = 1.54184 Å). Space group P212121, a = 4.7555(2) Å, b = 14.7379(7) Å, c = 22.971(1) Å, *α* = *β* = *γ* = 90◦, V = 1609.95(12) Å3, Z = 4, Dcalcd = 1.371 mg/cm3; *μ* = 0.847 mm−1, F (000) = 704. The final R indicates *R* = 0.0484 (2682), w*R*2 = 0.1337 (3174). Crystallographic data of 1 have been deposited in the Cambridge Crystallographic Data Center, with deposition number 2072655. Copies of the data can be obtained, free of charge, on application to CCDC, 12 Union Road, Cambridge CB21EZ, U.K. (fax +44(0)-1233-336033; email: deposit@ccdc.cam.ac.uk).

#### *3.4. Maryer's Method*

As reported [40], compounds **3** and **4** (each for 1 mg) were separately dissolved in HCl (1 mL) and incubated for 24 h. The hydrolysate was dried and dissolved in acetone. Then NaHCO3 and FDAA were added to incubate for 1 h. After being cooled, the mixture was dissolved in 50% aqueous CH3CN to yield FDDA derivatives. The corresponding standard amino acids were treated with the same procedures. The FDAA derivates were analyzed by HPLC at 254 and 340 nm by comparing the retention times with those of standards.

#### *3.5. Induced CD (ICD) Experiment*

Compound **8** and dimolybdenum tetracetate [Mo2(OAc)4] were resolved in dried DMSO. Their CD spectra were recorded immediately. Then the ICD spectra were measured every 3 min until they were stationary. The inherent CD data of compound **8** was subtracted to provide its induced CD spectrum as described previously [41,42].

#### *3.6. Anti-Food Allergic Experiment*

The in vitro anti-food allergic experiment was conducted according to the reported method [43]. Briefly, IgE-sensitized RBL-2H3 cells were treated with tested compounds for 1 h. Then cells were stimulated with dinitrophenyl-bovine serum albumin. The bioactivities were quantified by measuring the fluorescence intensity of the hydrolyzed substrate in an Infinite M200PRO fluorometer (Tecan, Zurich, Switzerland). Phosphate-buffered saline (PBS) buffer and loratadine were used as negative and positive controls, respectively.

#### **4. Conclusions**

From the deep sea-derived fungus *Penicillium griseofulvum* MCCC 3A00225, 10 new and 26 known compounds were obtained. The structures of the new compounds were determined by extensive analysis of their NMR and HRESIMS spectra, the absolute configurations were confirmed by different methods including the single X-ray crystallography, Marfey's method, and ICD experiment etc. (−)-Cyclopenol (**13**) showed the strongest in vitro anti-food allergic activity with an IC50 value of 60.3 μM in IgE-mediated RBL-2H3 cells.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/md19040224/s1, Figures S1–S60: 1D and 2D NMR spectra of **1**−**10**.

**Author Contributions:** X.-W.Y. designed the project; C.-P.X. isolated all compounds. Q.L. and G.L. performed the bioactive experiments. Z.S. provided the fungus. C.-L.X. conducted fermentation. D.C. and L.-Z.L. performed the ICD and Marfey's methods. T.-H.Z. obtained NMR data. C.-P.X., L.-Z.L., and X.-W.Y. wrote the paper, while critical revision of the publication was performed by all authors. All authors have read and agreed to the published version of the manuscript.

**Funding:** The work was supported by grants from the National Natural Science Foundation of China (21877022), the COMRA program (DY135-B2-08), and the Xiamen Southern Oceanographic Center (17GYY002NF02).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

## *Article* **E**ff**ect of Drying Methods on Lutein Content and Recovery by Supercritical Extraction from the Microalga** *Muriellopsis* **sp. (MCH35) Cultivated in the Arid North of Chile**

#### **Mari Carmen Ruiz-Domínguez 1,\*, Paola Marticorena 2, Claudia Sepúlveda 2, Francisca Salinas 1, Pedro Cerezal 1 and Carlos Riquelme 2**

Received: 30 September 2020; Accepted: 14 October 2020; Published: 26 October 2020

**Abstract:** In this study, we determined the effect of drying on extraction kinetics, yield, and lutein content and recovery of the microalga *Muriellopsis* sp. (MCH35) using the supercritical fluid extraction (SFE) process. The strain was cultivated in an open-raceways reactor in the presence of seawater culture media and arid outdoor conditions in the north of Chile. Spray-drying (SD) and freeze-drying (FD) techniques were used for dehydrating the microalgal biomass. Extraction experiments were performed by using Box-Behnken designs, and the parameters were studied: pressure (30–50 MPa), temperature (40–70 ◦C), and co-solvent (0–30% ethanol), with a CO2 flow rate of 3.62 g/min for 60 min. Spline linear model was applied in the central point of the experimental design to obtain an overall extraction curve and to reveal extraction kinetics involved in the SFE process. A significant increase in all variables was observed when the level of ethanol (15–30% *v*/*v*) was increased. However, temperature and pressure were non-significant parameters in the SFE process. The FD method showed an increase in lutein content and recovery by 0.3–2.5-fold more than the SD method. Overall, *Muriellopsis* sp. (MCH35) is a potential candidate for cost-effective lutein production, especially in desert areas and for different biotechnological applications.

**Keywords:** microalgae; *Muriellopsis*; spray drying; freeze-drying; lutein; supercritical fluid extraction

#### **1. Introduction**

In the past few decades, demand for bioprospection of microorganisms isolated from harsh environmental conditions has been increased because of their diverse biotechnological applications. Among microorganisms, microalgae are the most diversified photosynthetic organisms with high adaptability to different environmental conditions [1]. They are mainly classified as *Cyanophyta* (cyanobacteria), *Rhodophyta* (red algae), *Chlorophyta* (green algae), and *Chromophyta* (brown algae) [2,3].

Microalgae are characterized as natural sources of bioactive molecules such as phycobiliproteins, polysaccharides, carotenoids, lipids, fatty acids, polyphenols, and vitamins [4,5]. These compounds exhibit health benefits such as antibacterial, antifungal, antioxidant, and anticancer activities that are essential in pharmacological, nutraceutical, food, and biotechnological development [5–7]. Carotenoids belong to the class of terpenoids and are derived from the 40-carbon polyene chain and

xanthophylls as their oxygenated derivatives in the presence of –OH groups (e.g., lutein), oxi-groups (e.g., canthaxanthin), or both (e.g., astaxanthin) [8–10]. Pure lutein is an orange-yellow, crystalline, and lipophilic solid whose chemical name is β, ε-carotene-3,3 -diol (C40H56O2). It is beneficial to human health due to its potential to ameliorate cardiovascular diseases [11], various types of cancer [12], and age-related macular degeneration [13] because of its antioxidant potential. The species known to accumulate carotenoids efficiently are *Chlorella* sp., *Chlamydomonas* sp., *Dunaliella* sp., *Muriellopsis* sp., and *Haematococcus* sp.

In this study, we used *Muriellopsis* sp. (MCH35), having the potential of producing carotenoids, especially lutein, as the main oxygenated carotenoid. This strain was isolated from an arid region of the north of Chile, as described previously by Marticorena et al. [14]. This area is located in Antofagasta, a well-known region for its high solar radiation, an environmental factor that favors microalgal growth [15]. Several studies have reported strategies to enhance microalgal carotenoid production using unfavorable environmental conditions such as nutrient deficiency, intense irradiation, and salinity to photo-bioreactors design [16–18]. These factors affect not only photosynthesis and productivity of cell biomass, but also pathways and cellular metabolism, and thus alter the cell composition [16]. The isolation and selection of microalgae is a prerequisite for the successful industrial production of biomass and beneficial compounds. Two factors are important in their successful industrial production: lutein content and biomass productivity. Other factors, such as the presence of cell wall or content of other carotenoids, can also be considered. A high lutein content in microalgae is necessary to achieve cost-effective and adequate amounts of extraction [19].

After the harvesting process, microalgal biomass pretreatment and extraction of bioactive compounds are most important because of their effect on bioactive molecule recovery. A drying process is the most common pretreatment method that increases the shelf life of naturally occurring bioactive biomass [20,21]. A dehydration process represents an essential step for reducing microbial growth, avoiding oxidative reactions, and improving bioactive compound extraction, although they are expensive [22–24]. However, these methods can alter the stability of labile bioactive compounds because of thermal breakdown [25]. Therefore, techniques such as spray-drying (SD) or freeze-drying (FD) that can adjust drying temperature have been recommended by researchers [23]. For the extractions of bioactive compounds, such as carotenoid from microalgae, many conventional methods, including maceration or soxhlet extraction, are used [19]. However, these methods are extensive and require a relatively huge amount of solvents, and thus are expensive and less eco-friendly [26]. Therefore, the use of green extraction technologies is increasing for the extraction of bioactive compounds. Green extraction technologies enhance extraction time, recovery, selectivity, and mass transfer with decreasing consumption of solvents. Supercritical fluid extraction (SFE), pressurized liquid extraction, microwave-assisted extraction, ultrasound-assisted extraction, and high-pressure homogenization are some examples of green extraction technologies [26].

The aim of this work is to study the effect of two different methods of biomass drying on lutein content and recovery, extraction kinetics, and yield of *Muriellopsis* sp. extracted by using the SFE process. The microalga was cultured in an open-raceways reactor under adapted outdoor conditions in the arid north of Chile. After harvesting of cells, SD and FD methods were employed for removing water content from the microalgal biomass and for knowing an individual carotenoid benchmark profile by using conventional methods. In the case of SFE, the parameters such as temperature (40–70 ◦C), pressure (30–50 MPa), and percentage of co-solvent (0–30% ethanol) were studied on the basis of Box-Behnken designs. In a similar way, the Spline linear model was used to control parameters for optimal lutein recovery from *Muriellopsis* sp. (MCH35). Drying pretreatment and optimal lutein extraction parameters for the microalgal biomass were studied with a focus on using the strain for future biotechnological applications and minimizing production cost when grown on a pilot-scale in a harsh environment.

#### **2. Results and Discussion**

#### *2.1. Growth Parameters and Carotenoid Profile of Muriellopsis sp. (MCH35)*

Photoautotrophic cultivation is a growing condition wherein light is an energy source, and carbon dioxide is an inorganic carbon source used to form chemical energy by using photosynthesis [27]. To evaluate the growth performance and parameters of *Muriellopsis* sp. (MCH35), batch tests were performed by using UMA5 culture medium [28]. The specific growth rate (μ) was found to be 0.085 d−1, as calculated by using Equation (1),

$$
\mu = \text{L} \text{n} \text{ (C)} \text{(Ci)} \text{/t} \tag{1}
$$

and is in accordance with previously reported data on the green microalga *Nannochloropsis gaditana* with similar environmental conditions, such as sufficient light and nutrient availability [29]. Therefore, the UMA5 culture medium is suitable for the cultivation of *Muriellopsis* sp. (MCH35) as it contains an adequate amount of nutrients required for the microalga. Nitrogen concentration and nitrogen:phosphorus ratio are widely recognized as determining factors for the growth and composition of microalgae [30,31]. An exponential increase was observed from day 0 to day 12 in biomass concentration starting with 0.44 g/L and reaching to the final concentration of 1.34 g/L (Figure 1A).

**Figure 1.** Evolution of (**A**) biomass concentration and volumetric productivity, and (**B**) productivity per area and photosynthetic efficiency of the strain *Muriellopsis* sp. (MCH35) in batch mode culture adapted under outdoor conditions in the arid north of Chile.

Chew et al. [32] reported the *Dunaliella* sp. biomass concentration of 1.5 g/L (in a volume of 3.4 L) at the laboratory scale. For *Phaeodactylum* sp., the biomass concentration of 1.38 g/L in 5 L medium was observed, while for *Chlorella vulgaris,* it was in the range of 0.6 to 1.08 g/L·d (in a volume of 1.5 L). Similar values were obtained for the first two strains, and the difference in biomass productivity of *Muriellopsis* sp. and *Chlorella vulgaris* can be due to the optimized laboratory conditions and differences in sizes of the two strains. The volumetric productivity of *Muriellopsis* sp. culture reached 75.73 mg/L·d on day 12, and the productivity per area was 16.81 g/m2·d (Figure 1A,B). High productivity was reported in the *Nannochloropis gaditana* cultures with 400 mg/L·d and incident irradiance of 1.100 μE/m2·s [33]. Our experiments on *Muriellopsis* sp. were carried out with an incident irradiance of 1.400 μE/m2·s, under outdoor conditions. The difference in productivities can be due to the different cultivation modes used, as a batch mode can affect productivity compared with semi-continuous mode. During the cultivation, it is essential to determine the variation in chlorophyll fluorescence (Fv/Fm) by focusing on the suitability of culture conditions. This parameter represents a measure of the quantum yield of PSII (photosystem II) and identifies any damage to the protein complex caused by photo-inhibition [34]. Figure 1B shows the values of photosynthetic efficiency ranging from 0.58 to 0.7 during the cultivation. An acceptable level of physiological acclimatization exhibited by *Muriellopsis* sp. was confirmed in our study (Figures 1B and 2). In optimal cultivation conditions, the productivity of *Muriellopsis* sp. was ~0.60 at a pH of 7.9–8.2 and temperature of 16.3–19.6 ◦C, when cultivated in a semi-continuous mode by using Arnon culture medium [35]. Del Campo et al. [36] determined the limiting growth conditions such as pH of 6–9 and the temperature of 33 ◦C that stimulated carotenogenesis in *Muriellopsis* sp.

**Figure 2.** Temperature and pH and evolution of *Muriellopsis* sp. (MCH35) in a batch mode culture adapted under outdoor conditions in the arid north of Chile.

The main carotenoids present in the SD and FD biomass are shown in Table 1. These data are supported by Figure 3, showing an HPLC chromatogram with a diode array detector from the FD microalga.

**Table 1.** Individual carotenoid profiles present in *Muriellopsis* sp. (MCH35) from two modes of dry biomass by the conventional method of extraction.

\* Benchmark extraction. Mean values in the same column followed by different letters (a-b) are significantly different (*p* < 0.05).

**Figure 3.** HPLC (High Performance Liquid Chromatography) chromatogram of individual carotenoids present in *Muriellopsis* sp. (MCH35), extracted under conventional extraction (freeze-dried (FD) biomass) and measured at 450 nm by a diode array detector.

The high individual carotenoid content was observed in FD biomass, except for astaxanthin that was 1.5 times higher for the SD method than the FD method. A similar drying effect was reported by Ryckebosch et al. [37] in which they evaluated the effect of the SD and FD methods on fresh biomass and the storage stability of lipids and carotenoids in the diatom *Phaeodactylum tricornutum.* Their study showed better results on carotenoid contents in the fresh and FD biomass of algae than that in SD biomass at 48 h (denominated as short-term storage). Lutein is the main carotenoid profile that *Muriellopsis* sp. shows [19,36,38]. Our results on lutein content (particularly lyophilized cells) were in a similar range with that reported previously (in range of 4.0 to 6.0 mg/g dry weight) [39,40]. Better lutein content was obtained by using both the drying methods than that reported by Molino et al. [41], who studied lutein production in different microalgae in a comparative manner. Our results were in a similar range as that reported by Del Campo et al. [8] and had an advantage that our cultures were produced at a large scale using seawater. This is relevant when developing massive cultures in areas with scarce water resources, such as desert areas (Antofagasta Region).

We found other carotenoids such as zeaxanthin, violaxanthin, astaxanthin, and β-carotene (can be seen in Figure 3) in low levels in *Muriellopsis* sp., similar to those reported by Del Campo et al. [36], with a total concentration of 5.10 ± 0.53 and 6.15 ± 0.63 mg/g for the SD and FD processes, respectively. Despite their low levels in *Muriellopsis* sp., their presence can contribute to pharmacological, nutraceutical, food, and biotechnological applications [7,42]. Lutein and zeaxanthin reduce age-related macular degeneration [43], while β-carotene prevents cataracts, skin diseases, and other illnesses like cancer [42,44]. Moreover, violaxanthin is a potential anti-photoaging agent acting against ultraviolet-B radiation (UV-B; λ 280–315 nm) [45], and astaxanthin is considered super vitamin E [46] for its stronger antioxidant activity (500 times more effective than α-tocopherol), preventing arteriosclerosis, coronary heart disease, and ischemic brain development [42,46]. Therefore, although it would be beneficial to optimize the extraction technology of lutein as suggested by Di Caprio et al. [47], our results provide preliminary information on the carotenoid profile of *Muriellopsis* sp. (MCH35).

#### *2.2. E*ff*ects of Drying Processes on the Extraction Yield of Muriellopsis sp. (MCH35) by SFE*

The experimental extraction conditions and results of the Box-Behnken designs from two-mode dry biomass by the SFE process are given in Table 2. The range of extraction yield (Y) was 0.11–7.84% (*w*/*w*) for the SD biomass and 0.76–6.05% (*w*/*w*) for the FD biomass. Table 2 shows the effect of co-solvent on extraction yield at all the conditions. The low levels of extracts were obtained without the addition of the modifier, such as 0.11–0.44% for the SD and 0.76–2.07% for the FD biomass.

**Table 2.** Extraction yields (Y) and lutein content and recovery by SFE from spray- and freeze-dried *Muriellopsis* sp. (MCH35) using Box-Behnken experimental design. The general parameters were biomass loading = 2.0 g, CO2 flow rate = 3.62 g/min, and extraction time = 60 min.

Acronyms: Temperature (T), Pressure (P), and Ethanol (co-solvent). Standard deviation was less than 5% in all operating conditions (SD ≤ 5%, *n* = 3).

Supplementary Table S1 shows the statistical data of both drying processes with a *p*-value in terms of the goodness of fit of the model. It can be seen that ethanol, as a co-solvent, was a significant variable in the extraction process for two drying methods of biomass. These data are reinforced with the mathematical equations obtained in the statistical section and are summarized for the significant factors and their interactions (ethanol). Equations (2) and (3) were developed for an approximate mathematical model to maximize the extraction yield from *Muriellopsis* sp. dehydrated using the SD (Equation (2)) and FD (Equation (3)) methods:

$$\text{Yield (\%, uy\%)}=0.88 + 0.49 \cdot \text{Ethanol} + 0.0001 \cdot \text{T} \cdot \text{Ethanol} - 0.001 \cdot \text{P} \cdot \text{Ethanol} + 0.0007 \cdot \text{Ethanol}^2 \tag{2}$$

$$\text{Yield (\%, u\#w)} = 7.85 + 0.10 \cdot \text{Ethanol} + 0.002 \cdot \text{T} \cdot \text{Ethanol} + 0.0001 \cdot \text{P} \cdot \text{Ethanol} - 0.002 \cdot \text{Ethanol}^2 \tag{3}$$

where Yield is extraction yield in %, *w*/*w*, Ethanol is co-solvent in % *v*/*v*, T is temperature in ◦C, and P is pressure in MPa. These regression equations mathematically approach a model to maximize the yield from the two biomass drying methods based on the experimental results obtained in this work. Figure 4 also shows the results of extraction yields obtained by using the Pareto chart and RSM (Response Surface Methodology) using conditions such as 40–70 ◦C temperature and 0–30% *v*/*v* ethanol (extractant) with the optimal pressure of 30 MPa (original ranging from 30–50 MPa) using the two

modes of dry biomass by SFE. Figure 4A,B represent the extraction yield of *Muriellopsis* sp. SD biomass, for which the optimum value was 6.40% *w*/*w* at 57.14 ◦C and 30 MPa pressure with 30% *v*/*v* ethanol as a co-solvent. These data are similar to the experimental conditions shown in Table 2 (run 11) with a value of 7.84% (55 ◦C, 30 MPa, and 30% ethanol). Figure 4C,D shows the significant factors obtained by the Pareto charts and the extraction yield obtained by RSM from the FD biomass. Its optimum value was similar to that of SD biomass with 5.96% at 59.67 ◦C temperature and 30 MPa pressure with 30% *v*/*v* ethanol as a co-solvent. In this case, run 26 (see Table 2) was the experimental condition closest to the optimum value, with 6.05% *w*/*w*. In the SFE process, the highest extraction yield and recovery of bioactive compounds can be achieved by optimizing some critical parameters. Indeed, many studies have shown an ability to modulate CO2 polarity by using co-solvents such as ethanol, and thereby increasing the extraction yields [48,49]. Our results showed an increase in the extraction yields using maximum extractant volume (30% *v*/*v*), similar to other studies. However, no significant interaction was found between temperature and pressure for the extraction yield in both the drying methods. Conversely, other reports accentuate the effect of factors including pressure or temperature on the extraction yield. Mehariya et al. [50] evaluated the pressure factor for measuring lutein extraction by the SFE process from the green microalga *Scenedesmus almeriensis.* They reported an increase in its yield with an increase in pressure from 25 to 55 MPa at 50 ◦C and above 65 ◦C (CO2 flow rate of 7.24 g/min).

**Figure 4.** Pareto Charts and response surface curves of the combined effects of temperature (40–70 ◦C), pressure (30–50 MPa), and ethanol as co-solvent (0–30% *v*/*v*) on extraction yield from (**A**,**B**) spray- and (**C**,**D**) freeze-dried biomass of *Muriellopsis* sp. (MCH35), respectively. The ± signs are interpreted in the Pareto graph according to the area of significance of factor or interaction in the experimental design and Response surface curves were drawn at 30 MPa as optimal pressure.

Generally, an improving trend in the yield was observed in the case of FD biomass. In spite of various alternatives for drying methods available, both SD and FD are the most commonly used drying methods for high-value products [51]. In particular, FD is the best method for water removal and for the conservation of biochemical composition, although it is the most expensive process [51,52]. However, the SD process is faster than other methods, but it can damage different thermolabile components [37]. Therefore, our results showed that the solid-state of water during FD could protect the primary structure of the microalgal biomass with minimal reduction in volume and improving its general extraction yield.

#### *2.3. The E*ff*ect of Drying Methods on Lutein Recovery of Muriellopsis sp. (MCH35) by SFE*

SFE experiments based on Box-Behnken design were also performed to evaluate lutein content and recovery from *Muriellopsis* sp. (MCH35) using both drying processes. The SFE study focused on lutein as the main carotenoid as other individual carotenoids were in low levels. The effects of drying methods on lutein recovery are shown in Table 2 and complemented with Supplementary Material (data shown in Table S1). The solvent factor was the most significant in both drying processes. Indeed, the interaction was stronger in the SD process as a quadratic factor. Both drying processes showed almost similar lutein content, with slightly more lutein content in FD biomass. FD biomass showed the highest lutein content (60.69 mg/g extract) during run 18 at 40 ◦C temperature, 50 MPa pressure, and 15% ethanol, followed by run 27 at 55 ◦C temperature, 50 MPa pressure, and 30% ethanol (54.48 mg/g extract). However, SD biomass showed lutein content of 47.14 mg/g extract (run 14) under 55 ◦C, 40 MPa pressure, and 15%, followed by run 1 (45.47 mg/g extract) at 40 ◦C temperature, 30 MPa pressure, and 15% ethanol. High pressure caused an increase in lutein content in FD biomass, contrary to SD biomass. In both processes, the temperature employed was between 40 and 55 ◦C and the amount of ethanol ranged from 15% to 30%, *v*/*v*. It is evident that the main difference in the lutein yield was due to the drying methods. Particularly, high pressure causes a major variation in solvent density so that it can have a good extraction yield for lutein. Moreover, the amount of free water present in *Muriellopsis* sp. (MCH35) can affect lutein extraction because of the difference in the degree of dehydration between SD and FD methods [52].

In accordance with the above results, lutein recovery obtained by using SD and FD methods is given in Table 2. Figure 5 shows the results obtained by Pareto charts and RSM curves for the combined effects of temperature (40–70 ◦C), pressure (30–50 MPa), and ethanol as a co-solvent (0–30% *v*/*v*) on lutein recovery from SD (Figure 5A,B) and FD biomass (Figure 5C,D). The presence of ethanol was significant in both methods. A similar trend was observed in the SD method because, again, the quadratic ethanol factor was relevant in lutein recovery (can be confirmed by Supplementary Material such as Table S1). The mathematical models are included in Equations (4) and (5) for the optimization of lutein recovery from SD and FD microalga, respectively. The equations summarize factors that were significant (ethanol as co-solvent), including their interactions.

$$\text{Lutein recovery} \left( \%, u \psi u \right) = 32.06 + 2.48 \cdot \text{Ethanol} + 0.0002 \cdot \text{T} \cdot \text{Ethanol} - 0.002 \cdot \text{P} \cdot \text{Ethanol} - 0.04 \cdot \text{Ethanol}^2 \tag{4}$$

$$\text{Latstein recovery} \left( \%, \, u \langle w \rangle \right) = 8.06 + 0.43 \cdot \text{Etilanol} + 0.019 \cdot \text{T} \cdot \text{Etilanol} + 0.002 \cdot \text{P} \cdot \text{Etilanol} - 0.03 \cdot \text{Etilanol}^2 \tag{5}$$

where Lutein recovery is measured as %, *w*/*w*, Ethanol is co-solvent in % *v*/*v*, T is temperature in ◦C, and P is pressure in MPa. These regression equations again approach a mathematical model obtained by the statistical software to maximize lutein recovery from both drying methods based on the experimental results. The highest lutein recovery obtained in this study was 49.84–74.52% *w*/*w* for the FD biomass in the presence of ethanol (15–30%, *v*/*v*). Conversely, the pressure and temperature factors were variable and hence were non-significant in the process. Moreover, this set of experimental factors also obtained the highest extraction yield. The optimal factors determined by the statistical analysis for the FD biomass were similar to those cited above. However, optimal lutein recovery was lower than the results mentioned above for the FD biomass, such as 60.47% *w*/*w* at 60 ◦C, 50 MPa, and 29.9% ethanol (*v*/*v*), approximately. On the other hand, the SD method achieved poor lutein recovery compared to that obtained by the lyophilization method. Thus, we obtained the best experimental result between 23.25% and 30.23% *w*/*w* in the presence of ethanol and at intermediate to high temperatures. A low-pressure trend was defined by the SD biomass in the presence of other optimal conditions (58.14 ◦C, 30 MPa, and 25.5% ethanol), with a value of 28.24% lutein recovery (*w*/*w*).

**Figure 5.** Pareto Charts and response surface curves of the combined effects of temperature (40–70 ◦C), pressure (30–50 MPa), and ethanol as a co-solvent (0–30% *v*/*v*) on lutein recovery from (**A**,**B**) spray- and (**C**,**D**) freeze-dried biomass of *Muriellopsis* sp. (MCH35), respectively. The ± signs are interpreted in the Pareto graph according to the area of significance of factor or interaction in the experimental design and Response surface curves were made at 30 MPa for spray-dried and 50 MPa for freeze-dried microalga as an optimal pressure.

Other groups of scientists have also studied the effect of supercritical fluid parameters such as pressure, temperature, solvent, and CO2 flow rate on lutein recovery and purity from other sources. Mehariya et al. [50] investigated the effect of pressure (25–55 MPa), temperature (50 and 65 ◦C), and CO2 flow rate (7.24 and 14.48 g/min) on the green microalga *Scenedesmus almeriensis*. Their results showed improvements in the lutein recovery (~98%) and purity (~34%) with an increase in temperature, pressure, and CO2 flow rate. Yen et al. [53] reported 76.7% lutein recovery from *Scenedesmus* sp. in the presence of 70 ◦C, 40 MPa, and ethanol (30 mol%), which is in the similar range of our results. Wu et al. [54] extracted 87.0% of lutein from *Chlorella pyrenoidosa* by SFE in 4 h in the presence of 50 ◦C, 25 MPa, and modified CO2 with 50% ethanol. The factors optimized for lutein recovery from microalgae by the SFE process in our study are similar to those reported in previous studies. The effect of water content on the *Muriellopsis* biomass for lutein recovery has been evidenced in our SFE results. Some studies have reported that the optimal parameters for SFE can be obtained from samples that exhibit 3% to 12% of water content [55–57]. The presence of moisture in samples can act as a barrier for the diffusion of supercritical CO2 and extracted compounds [57]. However, other studies showed that the presence of water can enhance extraction kinetics and yields from plants and microalgae such as *Nannochloropsis oculata* [58,59], and water can play the role of co-solvent for polar compounds according to the type of matrix. We got the results emphasizing that optimal supercritical extraction was obtained in the samples with low moisture (lyophilized cells). Although FD is an expensive process for dehydration, an increase of lutein recuperation was significant. This can coalesce with growing the endogenous *Muriellopsis* sp. in seawater and under arid outdoor conditions to counteract the high costs of the dehydration process. Therefore, *Muriellopsis* sp. (MCH35) can be used as an efficient lutein producer for biotechnological applications, especially in desert areas.

#### *2.4. Global Yield and Kinetic Curve of Muriellopsis sp. (MCH35)*

Based on the yield results obtained for SFE by using Box-Behnken designs from the two modes of dry biomass of *Muriellopsis* sp., the condition selected for kinetic study was the central point of the experimental design, that is, 55 ◦C, 40 MPa, and CO2 + ethanol (85:15 *v*/*v*) flow rate, since temperature and pressure were non-significant (*p* < 0.05).

Figure 6A,B show overall extraction curves (OEC) of SD and FD biomass. A recovery of 2.54% and 5.10% of extracts was obtained after 150 and 160 min of extraction for SD and FD biomass, respectively. Similar weights (~2.0 g) of dry biomass were used for extraction, indicating that a double amount of extract was obtained by FD compared with that of SD. This gives an advantage to the biomass obtained by FD over SD since the resulting final powder is better in quality and quantity. This is due to the fact that drying by sublimation, as in the case of the FD technique, is better.

(**B**)

**Figure 6.** Overall extraction curves (OECs) of the experimental (♦) and predicted data for Spline (···· CER, —- FER, — DC) models, and recovery (), at P = 40 MPa, T = 55 ◦C, CO2 + ethanol (85:15 *v*/*v*) flow rate (3.305 g/min), for Spray-Drying biomass (**A**), and Freeze-Drying biomass (**B**). Abbreviations: constant extraction rate period (CER); falling extraction rate period (FER); diffusion-controlled rate period (DC); Pressure (P) and Temperature (T).

The OEC plotted for SD and FD biomass followed the SFE kinetics that were established by Meireles [60] and Jesus et al. [61]. The extraction process began with the CER period, characterized by the removal of easily extractable compounds by solvent and co-solvent, which was mainly controlled by the convective mass transfer in the fluid film around the powder particles. Following the CER period, the transition period began with a reduced extraction rate, wherein the extraction rate was controlled by mass transfer mechanisms through both convection and diffusion. This period is commonly called the FER period. When easily accessible solute became scarce in the microalgae matrix, intra-particle diffusion became the main mass transfer mechanism during SFE, and hence the OEC assumed a typical shape of diffusion curve with reduced extraction rate.

From the fitted data by the Spline linear model in Figure 6A,B, the OEC parameters were estimated as given in Table 3. The calculated tCER were 12.54 and 11.24 min with the accumulated extracts of 1.66% and 3.25%, and the recovery of 65.85% and 64.72% for SD and FD biomass, respectively. This is in agreement with a previous study that reported the recovery between 50% and 90% in the CER period [62]. Although tCER of SD differs by less than 1 min and about 1.13% of the recovery compared to FD, the accumulated extract in FD was double that in SD. The calculated tFER was 29.01 and 44.11 min for the accumulated extract of 2.01% and 4.15%, and the total recovery of 79.63% and 82.70% for SD and FD biomass, respectively. At this stage, the difference between tFER of SD and FD was ~15 min, and the accumulated extract of FD was more than double that for SD. In our research, a recovery > 75% was achieved in the FER period. The MCER and MFER values represent the extraction rate of the CER and FER periods respectively [61], with values of 0.0026 and 4.6 × 10−4 g/min for SD, and 0.0057 and 5.3 × 10−4 g/min for FD. Therefore, the extraction rate that produced CO2 + Ethanol flow in the FD biomass was more than double that for SD. These values of the extraction rates were lower than the values reported for MCER in peach almond oil (0.0084–0.0752 g/min) [63] and MCER and MFER of chañar almond oil (0.066–0.0124 g/min) [64].

**Table 3.** Adjusted parameters of the spline linear model to SFE from *Muriellopsis* sp. biomass at 55 ◦C, 40 MPa, and CO2 + ethanol (85:15 *v*/*v*) flow rate.

*b*0: Linear coefficient of the first line (CER); *a*1, *a*2 and *a*3 : Slopes of the lines 1, 2, and 3 corresponding to the periods CER, FER, and DC, respectively; *tCER*, and *tFER* : Times in the intercepts of the lines 1 and 2 and the lines 2 and 3, respectively; *mEXT*(*t*) : mass of the extract at time t in each period; *Yt*: Variable response of the sixth, and the seventh row, for the considered stage (CER, FER, and DC).

On the other hand, Y values (mg extract/g biomass) in the CER and FER periods of FD showed a similar trend as above, since they were 1.9 and 2.5 times higher than the SD values. Y value (g extract/g CO2 + ethanol 85:15 *v*/*v*) represents the extract ratio in the supercritical phase at the bed outlet and were 2.3 and 1.3 times higher for the CER and FER periods respectively, in the FD biomass than in SD.

In the final stage of the process or DC period, the difference between the FD and SD biomass was observed. Table 3 shows the coefficients of determination (R2) for all periods, such as CER, FER, and DC obtained by means of the Spline linear model and their corresponding coefficients, such as bo, a1, a2, and a3. The CO2 + ethanol (85:15 *v*/*v*) flow rate was 3.305 g/min and solvent to feed ratio (S/F) of 8.2 to 247.1 (5–150 min) and 8.2 to 260.5 (5–160 min) for SD and FD, respectively. In different studies performed by using the SFE-CO2 technique, Sanzo et al. [65] reported the lutein recovery of ~47% from the *Haematococcus pluvialis* dry biomass with the CO2 flow rate of 3.62 g/min, 50 ◦C temperature, and 40 MPa pressure in 120 min. Yen et al. [53] reported the lutein recovery of 76.65% from the *Scenedesmus* sp. dry biomass with the CO2 flow rate of 1.45 g/min, 30% ethanol, 47.5 ◦C temperature, and 40 MPa pressure in 60 min.