Currently, the potential for marine natural products as drug candidates has been recognized all over the world, and the field is constantly growing and developing. Additionally, the development of new chemical and physicochemical approaches and tools has led to the isolation and structure elucidation of novel minor marine secondary metabolites, which could not be isolated/detected in the past. The number of structures isolated each year has almost doubled over the past 20 years. This is illustrated by the fact that according to the report of John Faulkner, 869 new structures were isolated from marine organisms during the year 2000 [5]. A decade later, in 2010, the number had risen to 1003 substances per year, as reported by John Blunt et al. [6], and a very recent report by Anthony Carroll et al. described 1490 new molecules isolated in the year 2017 [7]. Moreover, the development of new and improved organic synthesis methods made possible the synthesis of promising active compounds in the amounts required for further preclinical and clinical studies.

Tremendous progress in the clinical development of marine-derived drugs has been achieved over the past 20 years. For example, during this period, six out of nine currently used drugs of marine origin have been approved by their corresponding authorities [8]. Focusing specifically on anti-cancer drugs, two more antineoplastic agents, Plitidepsin [9] and Polatuzumab vedotin [10], have been approved since we reviewed this topic in 2017 [11]. Hence, the full list of the marine-derived drugs used for cancer treatment at the end of 2019 included:

By the end of 2019, there were an additional 28 drug candidates in different stages of clinical trials. The majority of them (24 substances, 85%) are undergoing clinical trials as anti-cancer drugs. Interestingly, 20 out of 24 molecules (83%) are antibody-drug conjugates (ADC) and therefore consist of a small cytotoxic "warhead" molecule and a specific antibody targeting different cellular membrane proteins.

Currently, five compounds are undergoing testing in phase III clinical trials and are at the final step before approval can be obtained. Apart from the well-known Tetrodotoxin (HalneuronTM, *Wex Pharmaceutical Inc*.), which is being developed as a painkiller for chronic, severe to moderate cancer-related pain, the other four candidates are all investigated as potential chemotherapeutic agents. In more detail, these substances are the nectin-4 targeting ADC Enfortumab Vedotin (ASG-22ME, *Seattle Genetics*) which is being tested in several urogenital tumors; the synthetic analog of the fungian diketopiperazine Plinabulin (NPI-2358, *BeyondSpring Pharmaceuticals Inc.*), which is able to inhibit tubulin polymerization and is tested in brain tumors and non-small cell lung cancer (NSCLC); the DNA minor groove binder Lurbinectedin (PM01183, Zepsyre®, *Pharmamar*), a synthetic analog of trabectedin which currently is tested in small cell lung cancer (SCLC), ovarian and breast cancers; and the bacterial proteasome inhibitor β-lactone-γ-lactam Marizomib (NPI-0052, Salinosporamide A, *Triphase*), which has been examined in melanomas, lymphomas, NSCLC, pancreatic cancers, as well as multiple myelomas.

On the other hand, most of the molecules undergoing phase I and phase II trials are monomethyl auristatin E or F (MMAE or MMAF) conjugates with different antibodies [8]. It seems a safe bet that we may expect several MMAE/MMAF based drugs to make it into clinics in the next decade. Excitingly, there are around 2000 marine-derived molecules that show interesting *in vivo* biological activity, and more than 10,000 compounds isolated from marine organisms for which *in vitro* activity has been described [12].

Drug development is a very dynamic field, as new and old drug candidates regularly enter phase I trials for different cancers, go on to the next trial phases, or fail. Therefore, this information should be regularly revised. For those who are interested in the most up-to-date status of different drug candidates, we recommend visiting the Marine Pharmacology web-site (https://www.midwestern.edu/ departments/marinepharmacology.xml) ran and regularly updated by former Editor-in-Chief of Marine drugs Prof. Alejandro M. S. Mayer (Midwestern University, IL, USA), and his team. Vital information can also be obtained from www.clinicaltrials.gov, www.accessdata.fda.gov, or directly from the web-pages of pharmaceutical companies involved in marine drug development.

The main obstacle for novel marine-derived molecules on their way to becoming clinically useful drugs (i.e., anti-cancer drugs) is the generation of larger amounts of the active substance. At the beginning of the era of clinical use of marine drugs, naturally harvested or maricultured organisms were used as the main source of an active compound. As an example, the first hundreds of milligrams of halichondrin B and trabectedin were produced from marine invertebrates cultured in marine farms by the US National Cancer Institute [13], and PharmaMar [14], without which preclinical and clinical trials and further development of the meanwhile approved drugs Halaven® and Yondelis® would not have been possible. Nowadays, interesting compounds are largely produced by chemical synthesis or by modifying natural compounds that are mass-produced by microorganisms, either in wild-type or genetically modified cultures. However, macromolecules, such as marine polysaccharides and their derivatives (mainly used as food, as food additives, or in the cosmetic industry), are isolated and purified from natural sources like marine alga. There are some exceptions to this rule, e.g., several small molecules possessing anti-cancer activity isolated directly from the sponge *Aplysina aerophoba* marketed by BromMarin [15].

*Marine Drugs* (ISSN 1660-3397) is a leading journal on the research, development, and production of biologically and therapeutically active compounds from the sea. To document the dynamic field of marine anti-cancer pharmacology, the Topical Collection "Marine Compounds and Cancer" (http://www.mdpi.com/journal/marinedrugs/special\_issues/marine-compounds-cancer) was started three years ago [11]. This Topical Collection covers the whole scope from agents with cancer-preventive activity to novel and previously characterized compounds with anti-cancer activity, both in vitro and in vivo, and the latest status of clinical development from drug trials. Of note, compounds possessing pro-carcinogenic activity or mediating cancer cell survival are also within the scope of the collection. Owing to the importance of trial execution, special attention is given to current shortfalls and possible strategies to overcome obstacles in the area of marine anti-cancer drug development.

These are exciting times for scientists and physicians involved in the investigation and development of marine drugs! We invite the authors to share with us and all our readers your latest and forward-looking discoveries!

Dr. Sergey A. Dyshlovoy and Dr. Friedemann Honecker, Guest Editors, Topical collection "Marine Compounds and Cancer".

### **References**

© 2019 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 (http://creativecommons.org/licenses/by/4.0/).

### *Review* **Design and Synthesis of Anti-Cancer Chimera Molecules Based on Marine Natural Products**

### **Min Woo Ha, Bo Reum Song, Hye Jin Chung and Seung-Mann Paek \***

College of Pharmacy and Research Institute of Pharmaceutical Sciences, Gyeongsang National University, Jinju Daero 501, Jinju 52828, Gyeongnam, Korea

**\*** Correspondence<EMAIL_ADDRESS>Tel.: +82-55-772-2424

Received: 23 July 2019; Accepted: 16 August 2019; Published: 27 August 2019

**Abstract:** In this paper, the chemical conjugation of marine natural products with other bioactive molecules for developing an advanced anti-cancer agent is described. Structural complexity and the extraordinary biological features of marine natural products have led to tremendous research in isolation, structural elucidation, synthesis, and pharmacological evaluation. In addition, this basic scientific achievement has made it possible to hybridize two or more biologically important skeletons into a single compound. The hybridization strategy has been used to identify further opportunities to overcome certain limitations, such as structural complexity, scarcity problems, poor solubility, severe toxicity, and weak potency of marine natural products for advanced development in drug discovery. Further, well-designed marine chimera molecules can function as a platform for target discovery or degradation. In this review, the design, synthesis, and biological evaluation of recent marine chimera molecules are presented.

**Keywords:** chimera; chemical conjugation; marine natural products; anticancer agent; hybridization

### **1. Introduction**

The struggle for existence in a natural environment occasionally makes a species develop its own unique weapons such as speed, power, or even toxins. As these toxins possess biologically potent activity and unique modes of actions, these natural products have been regarded as a robust platform for further medicinal research [1,2]. Thus far, marine natural products have been particularly highlighted for their extraordinary bioactivity under highly diluted conditions [3–5]. Therefore, it is plausible to utilize marine natural products as a hit or lead compound in drug discovery and its further development [6,7].

Although marine natural products could have the powerful potential for drug discovery, there are also a few obstacles associated with them. First, it is rather difficult to secure a sufficient amount of these products for further study [8]. In most cases, the medicinal study of natural products requires a substantial number of test samples for elucidation of target protein/receptor and following signaling pathway. However, harsh conditions, difficulty in access, and scarcity of the target organism are hindrances to large scalable synthesis of important marine natural products, such as spongistatin 1 (13.8 mg from 400 kg of marine sponge) [9], phorboxazole A (95 mg from 236 g of *Phorbas* sp.) [10], discodermolide (7 mg from 434 g of *Discodermia dissoluta*) [11], bryostatin 1 (0.9~1.8 g from 10,000 gallons of *Bugula neritina*), [12,13] etc. The unmet needs from natural resources occasionally leads to samples being obtained via chemical synthesis, and it usually remains difficult to satisfy substantial supply requirements [14–16] (Figure 1).

**Figure 1.** Structure of certain bioactive marine natural products.

Second, structural complexity is another hurdle. The highly complex structure of marine natural products frequently makes it very difficult to modify or synthesize them on a large scale [17,18]. In order to improve the biological activity of these products, both chemical modification and related structure-activity relationship (SAR) study of marine natural products are necessary [19]. However, their highly complex structure hampers efficient modifications and any subsequent systematic research [20]. In order to address this structural complexity, a more simplified analog of marine natural products can be a breakthrough in drug discovery, as evident in the case of halichondrin B [21] and its simplified analog eribulin [22] (Figure 2). Although halichondrin B possesses extraordinary cytotoxicity against B-16 melanoma cells (IC50 0.093 ng/mL) and other tumor cell lines, its structure is too complex to be an advanced drug or drug candidate for related cancer therapy. Consequently, the truncated ketone analog eribulin of halichondrin B was developed through an entirely synthetic approach [23] and approved for treatment of metastatic breast cancer as a mesylate salt.

**Figure 2.** Halichondrin B [21] and its synthetic analog eribulin [22].

The third hurdle is biological activity and its selectivity. Since marine natural products are derived from marine organisms and not humans, their application to the human body may trigger undesired biological processes [24]. In order to solve this selectivity issue, various analogs are made and tested [25,26]. In addition, systematic study for mechanism of action or ligand-target binding usually follows valuable analogs [27]. Nonetheless, marine natural products continue to serve as a versatile starting point for drug discovery because of their unique structural framework and biological activity. Simultaneously, the strategy for the efficient modification of this complex molecule has also been evolving.

Molecular hybridization can be a good strategy for advanced marine natural products [28–31]. Certain drawbacks of marine natural products such as structural complexity or non-selective biological activities were overcome through a combination with other bioactive molecules. This combination strategy was evolved to develop chimera molecules, pursuing not merely a synergistic sequence but a systematically operating biological sequence. For example, the hybridization of a ligand and ubiquitin recruiting probe enables the degradation of target biomolecules via cellular sequence, such as ubiquitination and following the proteasome pathway. This proteolysis targeting chimera (PROTAC) approach showcases a well-designed chimera strategy from natural products [32–37]. (Figure 3) In addition, biotinylation of active ligand provides affinity column chromatography to enable target protein isolation [38]. In addition to these strategies, an improvement of basic activity can be anticipated. In this regard, advanced chimeric molecules from marine natural products with anticancer activity is reviewed here.

**Figure 3.** Comparison of the combination and chimera strategies.

### **2. Results**

### *2.1. Conjugation with Other Active Molecules*

The macrosphelide (MS) family [39], comprising 13 natural isomers, are 15- or 16-membered macrolide antitumor agents. They are derived from marine sponge *Periconia byssoides* [40] (MSC **1**, MSE-H, and MSL) and soil fungi *Microsphaeropsis* sp. FO-5050 [41] (MSA-D, MSJ, and MSK) and show cell adhesion inhibitory activity or immunosuppressive activity [42]. In addition, a derivatization of macrosphelide skeleton based on MSA **2** or MSE **3** enables the activation of apoptosis in human lymphoma U937 cells, although their activity is slightly weak [43]. In order to increase this apoptosis-inducing activity in cancer cells, preparation of a chimera compound with another 16-membered anticancer agent, epothilone **4**—which shows not only tubulin-disrupting profiles and anticancer activity, similar to the paclitaxel, but also an apoptosis-inducing property in cancer cells to inhibit tumor cell growth—was pursued [44]. Based on their similar structural features and biological properties, hybridization of MS and epothilone **4** was performed, pursuing an advanced anticancer agent.

Scheme 1 illustrates the synthetic plan of the MS-epothilone chimera. Known intermediate **5 [45]** for total synthesis of epothilone was prepared and linked to fragments **6**, **7**, [42] and **8** of MS via iterative esterification and deprotection of secondary alcohol moiety. For an elucidation of the structure-activity relationship, a thiazole side chain of epothilone was introduced in each CH3 group in the MS skeleton to provide the desired MS-epothilone chimera **9**–**11**.

**Scheme 1.** Hybridization of macrosphelide (MS) skeleton and epothilone side chain [44].

Screening of chimeras **9**–**11** exhibited an increase in their desired potencies, which is attributed to this hybridization strategy. In particular, chimera 9 showed more potent apoptosis-inducing activity in the U937 cell line as compared to other analogs or parent compounds MSA **2** or MSE **3**. It is noteworthy that other chimeras also showed an increased apoptotic property, while parent MSA **2** or MSE **3** did not have the same activity at the same concentration (1 μM, 12 h incubation). Further, successful hybridization of MS with epothilone **4** presents that the chimeric molecule strategy can be a powerful solution to the limitation of marine natural products themselves.

The complex structure of marine products can hamper efficient design or synthesis of advanced chimera molecules. In such cases, in silico study may offer another solution. Discodermolide **12**, a marine polyketide product from *Discodermia dissolute*, possesses potent antiproliferative activity against various human cancer cell lines [11]. Mechanically, it stabilizes microtubules and finally arrests cells in the mitosis status, just as paclitaxel does [46]. However, unlike paclitaxel **13**, a previous SAR and docking study of discodermolide at the paclitaxel binding site of tubulin revealed that free carbamate of discodermolide could be modified [47]. Based on the following study, conjugated diene in discodermolide resides in an aromatic pocket of tubulin. Moreover, the attachment of a simple aromatic group did not decrease its own anticancer activity. With this early study, the introduction of an aromatic side chain to discodermolide was pursued [47]. Synthetic intermediate **14** was coupled with various amine side chains, such as **15** with photolabile functionality, to finally produce carbamate **16** in good yield (Scheme 2). It was gratifying to note that the biological evaluation of this hybrid **16** showed an improved antiproliferative profile against human cancer cell lines. When it was treated to the lung cancer cell line A549, inhibition of cell growth at IC50 1.21 ± 0.35 nM was demonstrated, while discodermolide **12** showed a value of IC50 9.34 ± 0.56 nM (paclitaxel IC50 3.14 ± 0.09 nM). This pharmacological advance implies that the hybridization strategy based on the docking study could be another option for structurally complex molecules.

**Scheme 2.** Hybridization of discodermolide **12** with side chain moiety of paclitaxel **13** [47].

Further, hybridization of discodermolide **12** and dictyostatin **17** was also performed based on their similar structural features and anticancer activities, as depicted in Scheme 3 [48]. Dictyostatin **17** is a 22-membered macrolide natural product from the Indian [49] or Caribbean ocean sponge [50]. Dictyostatin has also garnered substantial attention due to its extraordinary antiproliferative effect (ED50 0.38 nM for P388 leukemia cell) [51]. A comparison of discodermolide and dictyostatin makes it plausible to say that they have similar backbones and substituents, except for the 22-membered lactone of dictyostatin. Based on this observation, hybridization of these two powerful anticancer natural products was performed [48]. During the synthesis and SAR study of discodermoilde **12**, a practical synthetic route to it was developed and applied to chimera preparation. *Para*-methoxybenzyl (PMP)-acetal **18** was coupled with aldehyde **19** using Wittig olefination to construct pivotal Z-alkene of target chimera. After some functional group interconversion and another Wittig olefination with diene **20**, discodermolide-dictyostatin chimera **21** was efficiently prepared. In addition, the biological activity of the chimera was also examined. Although chimera **21** possesses a relatively simple structure compared to discodermolide **12**, it displayed one-third the potency of discodermolide in a displacement test using [3H]-paclitaxel bound to microtubules. This simplified structure with moderate activity might open new possibilities for further development of related natural products.

**Scheme 3.** Structural comparison and hybridization of discodermolide **12** with dictyostatin **17** [48].

Since great cytotoxicity for solid tumors was observed using *Psammocinia* extracts from the waters of Papua New Guinea [52], significant effort has been made to isolate active ingredients produced from moderated cytotoxic marine natural products, such as cyclocinamide A [53], swinholide A [54], and furanosesterpenes [55]. However, these compounds did not explain the extraordinary cytotoxic properties of crude extract. The active ingredient was re-examined only in the early 2000s. Finally, highly anti-proliferative marine natural product psymberin **22** was isolated [52,56], but its absolute configuration and C4-stereochemistry were not confirmed. An additional research program elucidated its mysterious structure through total synthesis by the De Brabander group [57].

After confirmation of the correct structure, psymberin **22** was compared to classic natural compounds pederin **23** from rove beetles or mycalamide A **24** from marine invertibrates [58]. Although their structure possesses unique aminal-amide and trans-substituted tetrahydropyran skeletons, psymberin **22** has a characteristic dihydroisocoumarin moiety, while pederin/mycalamide A has another tetrahydropyran with the exomethylene group. As pederin/mycalamide A has been well reported as a powerful anticancer natural product based on eukaryotic protein synthesis inhibitors [52,56], a similar skeleton with a different side chain inspired the hybridization of the two natural products. It was anticipated that applying well-known biological properties of pederin to the psymberin-based would lead discovery [59] (Scheme 4).

**Scheme 4.** Hybridization of psymberin **22** with pederin **23** [59].

Bishomoallylic alcohol **25**, a synthetic intermediate for total synthesis of psymberin [57], was transformed into tetrahydropyranyl amide **26** in eight steps, with a 29% yield. It lacks a unique dihydroisocoumarin chain in psymberin, while it possesses a dimethoxy group in pederin. In order to introduce a side chain of psymberin, acyl halide **27** was attached and reduced with NaBH4. Final deprotection using LiOH/MeOH condition afforded separable C8-diastereomeric mixture of **28** and **29**. These two psymberin-pederin chimeras were used to unveil the SAR of this natural product family. It is interesting to note that the absence of dihydroisocoumarin moiety gave rise to a significant loss of psymberin's own cytotoxic property, while C8-epimer of the natural product caused a slight loss of this property. When they were treated to colon cancer cell line KM12, psymberin **25** or its C8-epimer showed a powerful or moderate toxicity (IC50 0.45 nM and 37 nM, respectively). However, its truncated analog **28** or C8-epi analog **29** showed little or no toxicity to the same cell line (IC50 710 nM and >1000 nM, respectively). These chimeras could be compared to pederin/mycalamide A as well. With a substituted tetrahydropyranyl side chain, which is rather common in natural pederin isomers, mycalamide A showed potent cytotoxicity (IC50 0.95 nM) as well. It was proven that the

common substituted side chain, or dihydroisocoumarin unit, plays a pivotal role in its own anticancer activity, employing a direct comparison of mother natural products and daughter chimeras. This case is a good indication of the effectiveness of a chimera strategy for drug development.

Topsentin **30** is a bisindolyl marine alkaloid isolated from several marine sponges in Mediterranean *Topsentia genitrix* or Korean *Spongosorites* sp [60]. As this alkaloid exhibits excellent cytotoxicity to cancer cells and features a rather simple structure when compared to the other active marine natural products [61], numerous researchers have attempted to develop more potent and drug-like analogs [62]. One of the indole side chains in topsentin was changed to an aminothiazole unit, as a thiadiazole in dendrodoine **31**, another marine alkaloid from *Dendroda grossular* [63]. With the first chimera diaminoindolylthiazole (DIT) **32** in hand, additional hybridization with curcumin **33** was performed to yield diaminocinnamoylthiazole (DCT) **34** and its analogs [64] (Scheme 5). Moreover, DITs and DCTs were screened to the cancer cell line. It is interesting that DITs are very effective in the induction of apoptosis in HeLa cells (IC50 1~45 μM), while DCTs play an active role in downregulating TNF-induced NF-κB activation. This research indicates that the hybridization of a simple functional group may also offer an opportunity to improve biological activity.

**Scheme 5.** Hybridization from topsentin **30**, dendrodoine **31**, and curcumin 1 **33** [64].

Neopeltolide **35** is another example of the chimera strategy of marine natural products. Since its isolation from marine sponge *Daedalopelta Sollas* in 2007 [65], urgent synthetic efforts were made to elucidate its correct structure [66]. As this marine macrolide showed various cytotoxic or cytostatic activity for numerous cancer cell lines, substantial research was conducted, including asymmetric synthesis, mechanism study, and analog synthesis. In addition, its antifungal activity was also focused upon. Contrary to its varied anticancer activity, its inhibitory activity to *Candida albicans* was highly potent at the minimum inhibitory concentration (MIC) 0.625 μg/mL [67]. This antifungal inhibition could be used for patients infected with the acquired immune deficiency syndrome (AIDS) or related fungal diseases. Further, the mechanism of action of neopeltolide was also studied to reveal that the agent works as the cytochrome bc1 complex inhibitor and inhibits the adenosine triphosphate (ATP) synthesis in mitochondria [68].

Although systemic research on neopeltolide was undertaken, its application in drug discovery was hampered by its complex structure. Neopeltolide **35** features a cis-substituted tetrahydropyran skeleton with an ansacyclic macrolactone framework. This formidable structure made it difficult to prepare neopeltolide on a large scale or related analogs in a varied manner. An interesting strategy was the chimeric application of neopeltolide with biaryl ether active molecules [69]. An SAR study reported that the oxazolyl carbamate skeleton plays an important role in its activity profile, while a complex macrolide skeleton has little effect on it [70]. Instead of this complex and less important left chain in neopeltolide, simple biaryl moiety was inspired by other similar fungicide metominostrobin **36**, famoxadone **37**, or other biaryl ethers, as illustrated in Scheme 6 [71].

**Scheme 6.** Hybridization from neopeltolide and biaryl ether hybrid [71].

Methyl propargyl carbamate **38** was converted into oxazolyl ester **39** in four steps and with high yield. Ester **39** was hydrolyzed and esterified with various primary alcohol **40** with biaryl moiety. Among synthesized chimeras, naphthyl benzyl ether **41** showed the most potent inhibitory activity (IC50 12 nM) for porcine succinate cytochrome c reductase (SCR). As this chimera simplified its mother structure, additional development for improved antifungal agent was expected.

Figure 4 presents the hybridization of iejimalide and archazolid skeletons [72]. Iejimalide B **42** and its family compounds were isolated from *Eudistoma cf. rigida* or *Cystodytes* sp., which were collected from an island in Japan [73]. Due to its remarkable anticancer activity and selectivity to NCI 60 cell lines, this polyunsaturated macrolide has been focused upon for its promising role in drug discovery [74]. However, its scarcity in natural resources led to not only total synthesis [75,76] but also structural simplification based on a hybridization strategy [72]. Archazolid A **43**, isolated from terrestrial myxobacteria [77], possesses a very similar structure as that of iejimalide B **42;** however, archazolid A **43** features a relatively simple side chain with thiazole and methyl carbamate moiety. In addition, because archazolid **43** showed potent inhibitory activity to vacuolar-type ATPases as well [77], iejimalide-archazolid chimera **44** was designed and synthesized.

**Figure 4.** Design of iejimalide B and archazolid chimeras [72].

Dibromothiazole **45** was converted to vinyliodide **46** using a 10-step sequence containing the Corey-Bakshi-Shibata (CBS) reduction [78], Marshall alkylation [79], and lithium-halogen exchange reactions as shown in Scheme 7. It was coupled with known catechol borane [76] **47**, using the Suzuki coupling condition [80] to afford tetraene **48**. Finally, the esterification of tetraene **48** with known carboxylic acid [76] **49** and ring closing metathesis [81] produced the desired chimera **44** in good yield. Biological evaluation of chimera **44** against various cancer cell lines was performed and it was proven that this hybridization yielded reduced toxicity to cancer cell lines, although certain cell lines such as lung adeno (LXFA 629L, IC50 0.32 μM) or colorectal (CSF HT29, IC50 0.49 μM)) were slightly sensitive to chimera **44**.

### *2.2. Conjugation with Other Functional Compounds*

For drug discovery and sequential development, marine natural products require more detailed studies, such as on-target protein isolation. The chemical conjugation of marine natural products with other functional compounds provides special opportunities in this regard. Biotin is a good example. Biotin, well known for its strong interaction with streptavidin [82–84], can be utilized to identify the target protein of natural products [85–88]. As this hybridization strategy may lead to the loss of its own binding affinity to natural products, it usually requires preliminary study to identify where to ligate biotin and what to use as a linker between biotin and the natural product. This pre-research requires additional efforts, thereby making it difficult to apply this method to a general natural product. However, in certain cases, well-designed chimeras showcase the usefulness of this hybridization protocol.

Bistramide A **52**, isolated from marine metabolite of *Lissoclinum bistratum* [89–91], is a good example of this hybridization. As the bistramide family has shown potent cytotoxic properties on various cancer cell lines, such as non-small cell broncho-pulmonary carcinoma or HL60 cells as well as unique spiroketal skeleton [92], this marine product has been a captivating target for synthetic chemists. After pioneering research on structural elucidation and synthesis of skeletons or other isomers [93], enantioselective total synthesis of bistramide A was accomplished in 2004 [94]. This synthetic route features iterative cross metathesis of terminal alkenes **53** and **55** with the geometrically strained cyclopropene **54** [95] in the presence of Grubbs second-generation catalyst **55** to construct bisunsaturated ketone **56**. The treatment of H2 with Pd(OH)2/C to this ketone induced hydrogenation/hydrogenolysis and spontaneous cyclization to afford the desired spiroketal **57** in 53% yield after the Dess–Martin oxidation sequence. Ketal **57** was converted to bistramide A **52** after employing amidation with other building blocks in six linear steps. This efficient and convergent synthesis made it possible to not only

elucidate the C37 chiral center [96] but also provide further opportunity for hybridization with other active molecules (Scheme 8).

**Scheme 8.** Total synthesis of bistramide A **52** [94].

Although potent and active, bistramide A **52** has been reported since its first isolation [97]; its mechanism of action was unknown until the preparation of bistramide-biotin chimera, as illustrated in Scheme 8 [98]. Its protein kinase C(PKC) δ inhibitory features caused PKCδ to be a cellular target in HL-60 cells. However, an in vitro study with real-time fluorogenic kinase assay system [99] revealed that bistramide A did not have strong affinity to PKCδ as its inhibitory activity. In order to identify an early-stage target, target protein fishing was planned. Based on the synthetic strategy previously described [94], homologation with carbon linker and final attachment of biotin was accomplished in order to produce desired chimera **58** as well as fragment-biotin chimera **59** for a controlled experiment (Figure 5). The treatment of these two chimeras into whole-cell lysate from A549 cell revealed direct binding of monomeric G-actin (K*<sup>d</sup>* 7nM) as a primary target protein. Using this target fishing study with biotin-chimera enabled the study of a more detailed mechanism of action.

**Figure 5.** Biotin chimera based on bistramide A [98].

Aplyronine A **60**, isolated from Japanese sea hare *Aplysia kurodai* [100], is another case for the conjugation of marine natural product with biotin to identify the interaction of actin. While bistramide A **52** binds to actin, aplyronine A binds to the actin/tubulin complex, which is a 1:1:1 trimeric complex [101]. The Kigoshi group treated aplyronine A with aqueous HCl to produce the corresponding aldehyde, which was condensed with hydroxylamine **61** to form an oxime bond as an inseparable mixture of aplyronine-biotin-diazine chimera **62** (Scheme 9). When it was treated to HeLa S3 cells, α/β tubulins appeared in SDS-PAGE with some other nonspecific binding proteins. In order to validate this Western blot result, purified actin (from rabbit) or tubulin (from porcine brain) was treated with chimera **62**. This validation study examined that unusual aplyronine/actin/tubulin heterotrimeric complex is responsible for its extraordinary antitumor effect through cell arrest at mitosis in HeLa S3 cells (IC50 0.45 nM) [102].

**Scheme 9.** Preparation of aplyronine-biotin-diazine chimera **62** [101].

Diazonamide A **63** is a marine natural product from *Diazona chinensis*, located on the ceilings of small caves on the Siquijor Islands in the Philippines [103]. Despite its scarcity in natural sources (54 mg from 256 g of *D. chinensis*), its extraordinary antimitotic property in human colon carcinoma and murine melanoma cancer cell lines has garnered substantial interest from chemists and biologists [104–106]. This tremendous study led to structural elucidation, efficient chemical synthesis, and related analog preparation. Its target protein study is interesting. When diazonamide (syndistatin **64**)-biotin chimera **65** was prepared and treated with HeLa extract, two additional bands of an affinity matrix were observed on an SDS/PAGE compared to those in the control experiment. Mass identification indicated that these two bands were ornithine δ-amino transferase, although why the matrix separates is unknown [107]. As ornithine δ-amino transferase is well known as a mitochondrial enzyme in the TCA cycle [108], it was slightly surprising. However, further study using the RNA*i*-mediated knockdown method and additional investigation proved that ornithine δ-amino transferase is essential for mitotic cell division. Thus, target protein and its corresponding pathways of diazonamide were unveiled to lead to the discovery of advanced diazonamide analog DZ2384 **66 [109]**. This process shows a 5–50 times higher efficacy without neurotoxicity at an effective dose (Figure 6).

**Figure 6.** Preparation of biotinylated diazonamide and its advanced analog DZ2384 **66** [107].

Further, macrosphelide (MS) skeleton was also utilized in this chimeric research, as depicted in Scheme 10 [110]. Although it has a structural framework and biological profiles, its mechanism of action is not well studied in the MS family. In order to identify the target protein of MSA **2** and elucidate the following pathway, MSA-biotin hybridization was planned. The protected Weinreb-amide of (*S*)-lactic acid **67 [111]** was transformed into allylic alcohol **68** using a three-step protocol—alkylnylatoin, carbonyl reduction, and alkyne reduction—in 42% yield. This key intermediate **68** in hand, allylation, iterative esterification, and deprotection sequence finally afforded allyl-MSA **69** in 7.6% yield and 13 steps. Biotin was linked to allyl-MSA **69** after additional manipulation. For efficient target fishing, a long carbon chain was selected as a linker. Terminal alkene in allyl-MSA was utilized in cross metathesis to introduce a long carbon linker and protect the amine functional group in amino MSA **71**. Finally, acidic deprotection and following amidation with biotin tag **73** produced the desired MSA-biotin chimera **74**. Target fishing studies, employing this chimera, on potent cell-cell adhesion inhibitor are ongoing.

**Scheme 10.** Preparation of biotinylated marcosphelide chimera **74** [110].

The marine natural product-biotin chimera strategy was also applied to identify the target protein of kahalalide F **75**. Ever since the isolation of kahalalide F from marine mollusk, *Elysia rufescens*, in 1993 [112], this polypeptide marine product has been used in clinical trials because of its powerful anticancer activity and low cellular toxicity [113]. For advanced development, identification of the target protein using biotinylation of kahalalide F was also attempted, as presented in Scheme 11. Kahalalide F was coupled with biotinylated linker **76 [114]**, prepared from biotin and tetraethyleneglycol, under a weak basic condition to produce desired chimera **77** in 85% yield. When chimera 77 was treated to T7 cDNA phage for reverse chemical proteomics, human ribosomal protein S25 was found to be responsible for its potent anticancer property in a dose-dependent manner.

**Scheme 11.** Preparation of Kahalalide F-biotin chimera **77** [114].

Avrainvillamide is another good example of the biotin chimera strategy [115]. In order to unveil the mysterious target protein of powerful avrainvillamide, isolated from *Aspergillus* sp. CNC358 [116], a biotinylated natural product **79** was designed. Synthesis of chimera **79** is depicted in Scheme 12. Iodoarene **80** [116] was stannylated to yield aromatic stannane **81** via metal/halogen exchange and a substitution reaction. Then, stannane **81** was coupled with vinyl iodide **82** [117] to produce nitroarene **83**, which was converted into biotinylated arene **85** employing cross metathesis with terminal alkene **84 [118]**. Reductive cyclization of **85** and HPLC purification afforded the desired chimera **79** (1:1 mixture of diastereomers at the C21 chiral center). Finally, Western blotting and MS/MS sequencing of these chimera-treated T-47 D cells revealed that nucleophosmin was a target protein of avrainvillamide. As nucleophosmin is a multifunctional protein in numerous tumors [119], a more detailed study was subsequently conducted to disclose the mode of action of avrainvillamide and its related natural product stephacidin [120].

**Scheme 12.** Preparation of avrainvillamide-biotin chimera **79** [115].

The hybridization strategy was also utilized to identify the target protein of pateamine, which is isolated from the New Zealand marine sponge [121]. As this marine natural product showed not only remarkable cytotoxicity to tumor cell line P338 [121] but also extraordinary immunosuppressive activity [122], target protein identification was pursued, as illustrated in Scheme 13 [123]. Primary iodide **88** [124] was alkylated with truncated pateamine **89** [123], which is more stable than pateamine **86**. Further, corresponding vinyl bromide **90** [124] was coupled with vinyl stannane **91** [122] to produce the desired pateamine-biotin chimera **87** in moderate yield. The pull-down method of this affinity-matrix **87** showed that it bound to and inhibited the association of eIF4A and eIF4B, finally inhibiting the cap-dependent eukaryotic translation sequence [124]. This fundamental study has also opened the way for more detailed research and further applications of pateamine analogs [125].

**Scheme 13.** Preparation of pateamine A-biotin chimera **87** [124].

### **3. Conclusions**

As an endless resource of bioactive molecules, marine natural products stand a reliable chance in the world of drug discovery. However, certain limitations of these products, such as complex structure, toxicity, selectivity, and even potency have driven researchers to identify additional strategies for these natural compounds. Chimeras yielded by the hybridization of two active molecules are one such solution. Changing the active skeleton of the mother framework also added more opportunities, but this process also comes with its own set of unique drawbacks, such as loss of function, scarcity of supply, and complex structure for chemical modification. In order to overcome these limitations, SAR, structural simplification, development of the synthetic route, and the medicinal chemistry approach have been studied thus far. Based on this fundamental progress, currently, hybridization of active natural products is being utilized to improve their own biological properties and elucidate veiled mechanism of actions. The efficient application of this chimera strategy to other active molecules remains a direction for groundbreaking research in the future.

**Funding:** This research was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF), funded by the Ministry of Education, Science and Technology (NRF-2016R1C1B2006699).

**Acknowledgments:** We are grateful to the two reviewers for their constructive input.

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

### **References**

© 2019 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 (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Selective Suppression of Cell Growth and Programmed Cell Death-Ligand 1 Expression in HT1080 Fibrosarcoma Cells by Low Molecular Weight Fucoidan Extract**

### **Kiichiro Teruya 1,2,\*, Yoshihiro Kusumoto 2, Hiroshi Eto 3, Noboru Nakamichi <sup>3</sup> and Sanetaka Shirahata <sup>1</sup>**

Received: 28 June 2019; Accepted: 14 July 2019; Published: 19 July 2019

**Abstract:** Low molecular weight fucoidan extract (LMF), prepared by an abalone glycosidase digestion of a crude fucoidan extracted from *Cladosiphon novae-caledoniae* Kylin, exhibits various biological activities, including anticancer effect. Various cancers express programmed cell death-ligand 1 (PD-L1), which is known to play a significant role in evasion of the host immune surveillance system. PD-L1 is also expressed in many types of normal cells for self-protection. Previous research has revealed that selective inhibition of PD-L1 expressed in cancer cells is critical for successful cancer eradication. In the present study, we analyzed whether LMF could regulate PD-L1 expression in HT1080 fibrosarcoma cells. Our results demonstrated that LMF suppressed PD-L1/PD-L2 expression and the growth of HT1080 cancer cells and had no effect on the growth of normal TIG-1 cells. Thus, LMF differentially regulates PD-L1 expression in normal and cancer cells and could serve as an alternative complementary agent for treatment of cancers with high PD-L1 expression.

**Keywords:** fucoidan; low molecular weight fucoidan extract; N-Ras; neuroblastoma-rat sarcoma; Cancer; programmed cell death-ligand 1; programmed cell death-ligand 2; human sarcoma cell line (HT1080 cells); human normal diploid fibroblast (TIG-1 cells)

### **1. Introduction**

Various types of cancers evade T-cell responses and host immunity via adaptive negative regulators or co-inhibitory (checkpoint) receptors, such as programmed cell death-1 (PD-1) and their respective ligands, programmed cell death-ligand 1 (PD-L1 or B7-H1), programmed cell death-ligand 2 (PD-L2), cytotoxic T-lymphocyte-associated antigen 4 (CTLA-4), and CD80 (B7.1) or CD86 (B7.2) [1,2].  In order to acquire cytocidal function, a variety of immune cells, including resting T cells expressing PD-1, must be activated by two stimulating signals from an antigen-presenting cell (APC), as shown in Figure 1 [3–5]. In addition, resting T-cells express T-cell receptor (TCR) and CD28 on their cell surface which interact with the antigen-specific signal presented on the major histocompatibility complex (MHC) and CD80 (B7.1) or CD86 (B7.2) ligand expressed on the surface of an APC, respectively. Interactions of these receptor-ligand pairs are referred to as the early stage of T-cell activation [1,4]. The activated T-cells express CTLA-4, and as the binding affinity of CTLA-4 for CD80/86 is 500- to 2500-fold greater than that for CD28, CTLA-4 (negative regulator) competes with co-stimulatory CD28 signals to bind to CD80/86 [6,7].

Conversely, the interaction of PD-1 expressed on the T-cell surface with its ligand PD-L1 or PD-L2 on APC causes inhibition of T-cell activation, leading to reduced cell proliferation, and induces T-cell cytolysis [6,8]. While a balanced PD-1/PD-L1 interaction is a prerequisite for maintaining the normal T-cell response and assuring normal cell survival, overexpressed PD-L1 on cancer cells in the tumor microenvironment react with the PD-1 expressed on T cells and induce T-cell apoptosis, thereby facilitating tumor survival, progression, and evasion of the host tumor immune surveillance system [4,7,9]. Moreover, higher PD-L1 expression in many tumors corresponds to enhanced immune evasion thereby promoting tumor growth by suppressing the T-cell response [7,10–12]. Therefore, considering the PD-1/PD-L1 axis, the selective inhibition of PD-L1 expressed on cancer cells is expected to maintain the host defense system and consequently contribute towards cancer eradication.

**Figure 1.** Simplified image of T-cell activation and cancer cell lysis. Resting T-cells are activated by the interaction of TCR:MHC and CD28:CD80 and CD86, leading to the expression of CTLA4. CTLA4 preferentially reacts with CD80 and CD86 causing activated T-cell lysis. Although imbalanced interaction of programmed cell death-1 (PD-1) on activated T-cell and programmed cell death-ligand 1 (PD-L1) on antigen-presenting cell (APC) causes T-cell lysis, such reaction between activated T-cells and cancer cells expressing PD-L1 will lead to the survival of cancer cells and facilitate cancer cell growth. Suppression of PD-L1 by low molecular weight fucoidan extract (LMF) leads to cancer cell lysis. Adapted from [7,13].

PD-L1 expression is upregulated not only by some inflammatory cytokines (e.g., interferon-γ) but also by oncogenic activation of signaling pathways [6,12]. In one of the two main oncogenic signaling pathways, the extracellular epidermal growth factor (EGF) and TGF-α stimulate the EGF receptor (EGFR) which in turn activates wild type rat sarcoma oncogene homolog (Ras) to activate the raf pathway [14]. Moreover, in the second pathway, the constitutively active mutant Ras activates its immediately downstream factors, including phosphatidylinositol 3-kinase (PI3K), Rac1, and Raf which trigger a sequential activation of multiple downstream factors and eventually induce expressions of many genes such as RhoA/B, EGFR, VEGF, and PD-L1 in cancer cells [2,6,15–17].

Other lines of evidence have shown constitutive activation of mutant N-*ras* allele encoded in a human soft tissue sarcoma cell line, HT1080, which regulates Raf and RhoA pathways. Moreover, Raf-activated MEK, an intermediary transducer, is essential in the Ras signaling pathway but not in the PI3K/Akt pathway that includes nuclear factor-kappa B (NF-κB), resulting in the aggressive tumorigenic phenotype in HT1080 cells [18,19]. These data suggest that the Raf regulated pathway is closely associated with carcinogenesis, implying that this pathway regulates PD-L1 expression as one of the terminal steps in HT1080 cells. However, these studies do not address the involvement of Ras-regulated pathways in PD-L1 expression in HT1080 cells. As the PD-1/PD-L1 axis plays a major role in human cancers for immune evasion [17], it might be extremely valuable to cancer patients if

prospective agents such as fucoidan devoid of side effects are available to regulate PD-L1 expression exclusively in cancer cells.

Fucoidan can be found mainly in various species of brown algae (brown seaweed) such as wakame (*Undaria pinnatifida*), kombu (*Saccharina japonica*), mozuku (*Cladosiphon okamuranus*), hijiki (*Sargassum fusiforme*), bladderwrack (*Fucus vesiculosus*), and hibamata (*Fucus evanescens*) [20,21]. Fucoidan is fucose containing sulfated polysaccharides exhibiting complicated chemical structures due to various combinations of fucose, uronic acids, galactose, and xylose. [22–24]. The structure of fucoidan from *Cladosiphon okamuranus* belonging to Genus *Cladosiphon* had been determined partly using the nuclear magnetic resonance (NMR) method [25] and we reported that this structure possess similar structural features as the fucoidan from *Cladosiphon novae-caledoniae* Kylin. It has been noted that Quantitative 1H-NMR (qNMR) analysis for the qualitative and quantitative characterization of metabolites may be applied for crude biological extracts such as fucoidan [26]. Just recently, the chemical structure of fucoidan from *Cladosiphon okamuranus* was determined using NMR analyses [27], as shown in Figure 2, and we now also refer to this structure as having a similar structural feature as fucoidan from *Cladosiphon novae-caledoniae* Kylin.

**Figure 2.** Structure of fucoidan from mozuku (*Cladosiphon okamuranus*) [27].

Fucoidan extracts usually show various sizes, such as 5100 kDa [28] and 1600 kDa [29], depending on the extraction methods and the kinds of seaweeds. Various molecular weight derivatives designated as low-molecular-weight fucoidan (LMWF: up to 40 kDa), intermediate-molecular-weight fucoidan (110–138 kDa), and high-molecular-weight fucoidan (300–330 kDa) [30], and other size classifications up to 10,000 kDa [31], have been prepared for biological and biochemical characterizations. In still other cases, many derivatives having molecular weights ranging between 7.6 kDa and 712 kDa have been assessed for their membrane permeability in vitro, and bioavailability in humans [32–36]. Apart from the above higher molecular weight derivatives, there are a group of fucoidan derivatives possessing much lower molecular weights designated as low molecular weight fucoidan (LMF) also called as oligo-fucoidan having MW of 500–800 Da [37], LMF from L. japonica (LMF-LJ) having an average molecular weight of <667 Da [38], low molecular weight fucoidan (LMWF) having main MW of 760 Da [39], and LMF having an average MW of 800 Da [40]. Finally, low molecular weight fucoidan extract (LMF) having a main MW of <500 Da, prepared by digesting crude fucoidan extracted from mozuku (*Cladosiphon novae-caledoniae* Kylin) with an abalone glycosidase, which was used in this study [41]. These two size groups of fucoidan derivatives have been examined for their health benefits as well as for their therapeutic effects and were shown to exhibit broad biological activities such as anti-tumor, antioxidant, anticoagulant, anti-inflammatory, and immune-modulatory effects in in vitro and in vivo studies [22,23,42–44]. With regards to its anti-tumor activity, fucoidan has been shown to exhibit suppressive effects in lung, breast, liver, colon, prostate, and bladder cancer cells [22,45]. In addition, our previous in vitro studies revealed that LMF (MW < 500 Da) can enhance the anticancer activity of chemotherapeutic agents (such as cisplatin, tamoxifen, and paclitaxel) [46] and it also demonstrated beneficial immunomodulatory effects in a clinical trial [23]. It has been shown that the low molecular weight fucoidan with a MW of 7.6 kDa exhibited higher intestinal absorptivity than the medium molecular weight fucoidan (MW 35 kDa) when assessed in the plasma and urine after oral administration in rats [36]. This result suggests that the lower molecular weight fucoidan derivatives are superior to the ones with higher MW in the intestinal absorptivity which is in line with our long-time notion of using LMF (mainly MW < 500 Da) for clinical application along with collecting supportive basic research data [23,46,47]. In order to further support these conclusions, the low molecular weight fucoidan derivatives mimicking the structural features of the genus *Cladosiphon* have been synthesized and tested for their anti-cancer activities. Results showed that one of the sulfated tetrafucoside synthetic derivatives could reduce MCF-7 and HeLa cell growth while showing no cytotoxic effect on normal WI-38 cells [48]. Moreover, it has been suggested that the most important factor that affects the biological activities of fucoidan is the branching degree of the fucoidan rather than its molecular weight, monosaccharides composition, and sulfate degrees [49]. Thus, it could be envisioned that an efficiently absorbed LMF (main MW < 500 Da) in the small intestine is perhaps also distributed throughout the intercellular environments, including tumor microenvironments, and such localized LMF may suppress cancer cell growth. These data together present the presently used LMF as an advantageous molecule over high molecular weight derivatives in further pursuing studies for basic research and clinical applications.

The molecular mechanism of fucoidan action involves suppression of the growth factor-stimulated pathway involving Ras/Raf cascade. A large amount of data supports the fucoidan inhibition of phosphorylation of intermediary transducers such as ERK1/2, PI3K, Akt, mTOR, c-Jun, c-Fos, EGF receptor, and vascular endothelial growth factor (VEGF) [21,22,39].

Accumulated data suggest that Ras regulates several downstream pathways to upregulate PD-L1 expression, while fucoidan suppresses Ras-regulated downstream signal transducers that coincide with those of the PD-L1 expression pathways. However, none of the preceding studies, as well as the relevant computer literature searches, explored the possibility that fucoidan suppresses PD-L1 expression regulated by the Ras-activated pathways. Therefore, we focused on examining the regulatory effects of LMF for several growth factors, PD-L1/PD-L2 and RhoA/B. We then examined our hypothesis that LMF differentially regulates PD-L1 expression in cancer and normal cells.

### **2. Results**

### *2.1. Suppression of Cancer Cell Growth by Low Molecular Weight Fucoidan Extract (LMF)*

To confirm whether LMF specifically suppresses cancer cell growth, HT1080 fibrosarcoma cells and normal human fetal lung diploid fibroblast TIG-1 cells were treated with varying concentrations (0–500 μg/mL) of LMF for 48 h. WST-1 cell viability assay showed significant dose-dependent suppression of HT1080 cell growth by LMF (\*\* *p* < 0.01, \*\*\* *p* < 0.001) (Figure 3A), while there was no statistically significant suppressive effect on TIG-1 cell growth up to 500 μg/mL (Figure 3B). Moreover, the response of HT1080 cells to concentrations lower than 100 μg/mL of LMF showed a similar dose-dependent suppressive effect with both WST-1 and Hoechst 33342 DNA staining assays (Figure 3C). In addition, Hoechst 33342 dye staining method was employed as this dye stains DNA of both live- and apoptotic-cells [50,51] resulting in the measurement of cumulative numbers of both cell types and allows the visualization of the cytocidal effect of LMF when compared with the WST-1

assay, which measures only live or metabolically active cell numbers [52,53]. The results showed a significant dose-dependent reduction in the cumulative cell numbers (Figure 3C). A comparison of IC50 of WST-1 as 40.1 μg/mL and IC50 of Hoechst staining as 73.7 μg/mL revealed that approximately 12% of HT1080 cells are apoptosis-induced cells by LMF. These results demonstrate that LMF at even lower concentrations suppresses cell growth dose-dependently, and approximately 12% of HT1080 cell apoptosis was observed to be induced by 36.6 μg/mL of LMF up to 100 μg/mL.

**Figure 3.** Effects of low molecular weight fucoidan extract (LMF) on cancer cell and normal cell growth. (**A**) HT1080 cells (1 <sup>×</sup> <sup>10</sup><sup>4</sup> cells/mL) and (**B**) TIG-1 cells (3 <sup>×</sup> <sup>10</sup><sup>4</sup> cells/mL), were plated (100 <sup>μ</sup>L per well) in a 96-well microplate. After 24 h, varying concentrations (0, 50, 100, 200, 500 μg/mL) of LMF were added, the cells were incubated for another 48 h, and live cell numbers in each well were measured by the WST-1 assay. The measured values were used for statistical analyses by Student's *t*-test. (**C**) Effect of lower LMF concentrations on HT1080 cell growth was examined using varying concentrations (0, 20, 40, 60, 80, 100 μg/mL) of LMF. After 48 h of treatment, cellular metabolic activity-based live cell numbers were measured by WST-1 (open circles, -), and live cell numbers were measured by Hoechst 33342 DNA staining (closed circles, -). The number of fucoidan untreated HT1080 cells was set to 100% to calculate the relative values of fucoidan-treated HT1080 cells. Statistical significance was determined by ANOVA with Tukey test. Different letters indicate statistically significant differences between each letter (*p* < 0.05). Half-maximal inhibitory concentration (IC50) values were calculated using approximation formulae.

### *2.2. Comparison of PD-L1 Expression Levels in Seven Cell Lines by Immuno-Fluorescent Staining*

Prior to examining the effect of LMF on PD-L1 expression, we compared PD-L1 expression levels in seven cell lines, using A549 and PC-9 cells as controls for low and high PD-L1 expression, respectively [12,44,54]. As shown in Figure 4, HT1080 cells expressed 1.06-fold higher PD-L1 compared to PC-9 cells. TIG-1 cells expressed PD-L1 levels comparable to those of the control cells (PC-9). Other cell lines expressed PD-L1 levels similar to or higher than those of the A549 cells. Therefore, we selected the HT1080 cell line for further studies to assess the effects of LMF on PD-L1 expression.

**Figure 4.** Comparison of programmed cell death-ligand 1 (PD-L1) expression levels in seven cell lines by immunofluorescence staining. Seven cell lines with indicated cell densities were prepared (HT1080 and A549 cells at 2 <sup>×</sup> <sup>10</sup><sup>4</sup> cells/mL, MCF-7, PC-9, NIH:OVCAR-3, and PANC-1 cells at 3 <sup>×</sup> 104 cells/mL, and TIG-1 cells at 4 <sup>×</sup> <sup>10</sup><sup>4</sup> cells/mL) and seeded at 100 <sup>μ</sup>L per well of a 96-well black plate followed by 24 h culture. PD-L1 detection was carried out using the anti-PD-L1 antibody, as described in the Methods section. Cells were stained with Hoechst 33342 for 30 min and analyzed using (**A**) IN Cell Analyzer 1000. The fluorescence intensities of each cell line were measured and converted to a numerical form for graphical presentation. For each cell line, six viewing fields per well were analyzed. A549 and PC-9 cells were used as low- and high-level controls for PD-L1 expression, respectively. (**B**) Four representative images acquired in (**A**) were shown. A, HT1080; B, A549; C, PANC-1; D, NIH:OVCAR-3.

*2.3. LMF Di*ff*erentially Regulates Programmed Cell Death-Ligand 1 (PD-L1) and PD-L2 mRNA Expression in HT1080 and TIG-1 Cells*

We examined LMF regulation of PD-L1/L2 expression in HT1080 and TIG-1 cell lines. HT1080 cells treated with 1 and 10 μg/mL of LMF for 48 h showed a significant reduction in PD-L1 (all, \*\* *p* < 0.01) and *PD-L2* (1 μg/mL, \* *p* < 0.05; 10 μg/mL, \*\* *p* < 0.01) mRNA levels (Figure 5A). TIG-1 cells treated with 10 and 100 μg/mL of LMF for 48 h showed significant increases in PD-L1 (10 μg/mL, \*\*\* *p* < 0.001; 100 μg/mL, \*\* *p* < 0.01) and PD-L2 (all: \*\*\*\* *p* < 0.0001) mRNA levels (Figure 5B).

**Figure 5.** Low molecular weight fucoidan extract (LMF) differentially effects transcription of programmed cell death-ligand 1 (PD-L1) and PD-L2 mRNAs in cancer and normal cells. (**A**) HT1080 and (**B**) TIG-1 cells were treated with varying concentrations (0, 1, 10 μg/mL and 0, 10, 100 μg/mL) of LMF respectively, for 48 h. After treatment, RNAs were isolated and subjected to quantitative reverse transcription polymerase chain reaction (qRT-PCR) analyses. Values obtained from LMF-treated cells were compared with those of untreated cells using Student's *t*-test (\* *p* < 0.05; \*\* *p* < 0.01, \*\*\* *p* < 0.001; \*\*\*\* *p* < 0.0001, *n* = 3).

### *2.4. Suppression of PD-L1 Protein Expression by LMF*

To further test the effects of LMF on PD-L1 expression, we measured the PD-L1 protein levels using an anti-PD-L1 antibody to test whether *PD-L1* mRNA suppression reflects similar suppression in protein levels. Western blot analysis of LMF-treated and untreated HT1080 cells was performed as described in the Methods section, and this method has been used by us to see the effects of fucoidan on HT1080 cells as well as other cultured cells [47]. PD-L1 protein band intensities of LMF-treated cells were lower than that of the control cells (Figure 6A). Protein bands in Figure 6A were quantitated for statistical evaluations and results show a significant reduction in PD-L1 protein (\*\*\* *p* < 0.001, Figure 6B) in LMF-treated HT1080 cells. Therefore, the PD-L1 protein levels reduced proportionally with a reduction in mRNA and the results suggest that the upstream signal transducers are mostly affected for PD-L1 transcription.

**Figure 6.** Low concentration of low molecular weight fucoidan extract (LMF) suppresses programmed cell death-ligand 1 (PD-L1) protein expression. (**A**) Western blot detection of PD-L1 protein in total protein lysates from HT1080 cells treated with 10 μg/mL of LMF for 5 days by replacing every 2 days with fresh media containing 10 μg/mL of LMF. Control untreated HT1080 cells treated similarly but without LMF. (**B**) Chemiluminescence protein bands from LMF-treated and untreated samples quantitated and bands from LMF-treated cells compared with that of untreated cells using Student's *t*-test. (\*\*\* *p* < 0.001, *n* = 3).

### *2.5. Suppression of Cell Surface PD-L1 Protein Expression by LMF*

Western blot analysis showed reduced PD-L1 protein levels in the HT1080 cell protein lysates proportionally with a reduction in PD-L1 mRNA levels. However, this method detects total PD-L1 proteins including intracellular premature PD-L1. The mature form of PD-L1 protein is anchored to the cell membrane with two extracellular domains [6]. It is necessary to show the level of surface PD-L1 exquisitely, as it is performing the most important and integral role in the immune escape mechanism of cancer cells. For this, we assessed if the effects of LMF are due to changes in the cell surface PD-L1 protein levels using the anti-PD-L1 antibody and analyzed this using flow cytometry, according to the published method. This method has previously been used to measure the PD-L1 expressed on the surface of several cell lines such as A549, H1975, and others using PD-L1 antibody followed by the flow cytometry analysis using a FACScan instrument [12]. Fluorescence histograms showed a small shift towards the left in LMF-treated HT1080 cells (Figure 7A, red line) compared with untreated control cells (Figure 7A, blue line), suggesting reduced levels of membrane-bound PD-L1 protein. To confirm this effect, mean fluorescence intensity (MFI) per cell, derived from histogram data, is presented in Figure 7B. These results show significant reduction in cell surface PD-L1 protein in LMF-treated HT1080 cells compared with the control cells (\*\* *p* < 0.01). Thus, the results show that the PD-L1 mRNA level corresponds with that of the total as well as cell surface PD-L1 protein levels, suggesting an effect on upstream signal transducers regulating PD-L1 transcription.

**Figure 7.** Effect of low amount low molecular weight fucoidan extract (LMF) treatment for 5 days on programmed cell death-ligand 1 (PD-L1) protein expression in HT1080 cells. HT1080 were treated with a low concentration of LMF (10 μg/mL) for 5 days and changes in PD-L1 protein levels were measured by flow cytometer. (**A**) PD-L1 protein expressed on the cell surface of HT1080 cells with (red line) and without (blue line) LMF treatment detected using anti-PD-L1 antibody and histograms were generated. (**B**) Mean fluorescence intensity (MFI) per cell was calculated using histogram data. Student's *t*-test (\*\* *p* < 0.01, *n* = 3) compares MFI of LMF-treated cells with that of untreated cells.

### *2.6. LMF Does not Interfere with the Binding of PD-1 with PD-L1*

The binding of PD-1 and PD-L1 occurs between T-cells and APCs for a normal T-cell response [3]. PD-1 binding with the PD-L1 expressed on cancer cells is an important step to evade the host immune surveillance system. Considering this, if LMF binds to PD-1 or PD-L1, attenuation and eventual inhibition of the normal T-cell response will occur. Therefore, we examined whether LMF inhibits binding of PD-1 with PD-L1. For this, we used PD-1 neutralizing antibody as a positive control which,

as expected, showed dose-dependent inhibition of PD-1 and PD-L1 binding (Figure 8A). When LMF replaced the positive control in the same experimental setup, it did not interfere with the PD-1 and PD-L1 binding (Figure 8B). Therefore, the results show that LMF does not interact with PD-1 or PD-L1 and suggest that its suppressive effect is due to regulation of the other cell surface proteins as well as intracellular transducers involved with expression of PD-L1 and other genes.

**Figure 8.** Inhibitory activity of programmed cell death-1 (PD-1): PD-L1 binding by low molecular weight fucoidan extract (LMF). (**A**) We used PD-1 neutralizing antibody as a positive control in dose-dependent inhibition of PD-1 and PD-L1 binding. Statistical significance was by ANOVA with Tukey test. The letters, a, b, and c indicate statistically significant differences between each letter (*p* < 0.05). (**B**) LMF was used to observe a dose-dependent inhibition of PD-1 and PD-L1 binding (N.S.: Not Significant).

### *2.7. LMF-Suppression of Epidermal Growth Factor Receptor (EGFR) and Vascular Endothelial Growth Factor (VEGF) Expression*

Growth factors play a significant role in carcinogenesis and metastasis [2,55]. The effect of LMF on EGFR, VEGF, and cell surface-associated protein expression was tested in HT1080 cells treated with different concentrations of LMF (0, 10, 100 μg/mL) for 24 h and mRNA levels were measured. LMF suppressed both EGFR mRNA (10 μg/mL, \* *p* < 0.05; 100 μg/mL, \*\* *p* < 0.01), and VEGF mRNA (10 μg/mL; \*\* *p* < 0.01, 100 μg/mL; \* *p* < 0.05) in HT1080 cells (Figure 9).

**Figure 9.** Suppression of epidermal growth factor receptor (EGFR) and vascular endothelial growth factor (VEGF) transcription by low molecular weight fucoidan extract (LMF). HT1080 cells were treated with 0, 10, and 100 μg/mL of LMF for 24 h. Total RNA was isolated and subjected to qRT-PCR analyses. LMF-treated cells were compared with untreated cells, and statistical evaluation was carried out using the Student's *t*-test. (**A**) EGFR, (**B**) VEGF. (\* *p* < 0.05, \*\* *p* < 0.01, *n* = 3).

#### *2.8. Suppression of RhoA and Stimulation of RhoB Expression by LMF*

The Ras superfamily contains Rho GTPases which are involved in many cellular processes that regulate various signal transduction cascades that affect gene expression, cell cycle progression, cell migration, and many other cellular events by cycling from GTP-bound active state to GDP-bound inactive state [56]. In addition to their fundamental functions, Rho GTPases such as RhoA and RhoB play a significant role in cancer progression and inflammation [57]. To assess the effect of LMF on RhoA and RhoB expression, HT1080 cells were treated with different concentrations of LMF (0, 10, 100 μg/mL) for 24 h and mRNA levels were measured. LMF suppression of RhoA gene transcription at 10 μg/mL (\*\* *p* < 0.001) and 100 μg/mL (\* *p* < 0.05) (Figure 10A) was significant. In contrast, fucoidan caused a significant (all, \*\**\* p* < 0.001) and dose-dependent (10 and 100 μg/mL) upregulation of RhoB gene transcription in HT1080 cells (Figure 10B). All the results obtained are summarized in Table 1.

**Figure 10.** Low molecular weight fucoidan extract (LMF) suppresses RhoA and stimulates RhoB transcription. HT1080 cells were treated with 0, 10, and 100 μg/mL of LMF for 24 h. Total RNAs were isolated and subjected to qRT-PCR analyses. LMF-treated cells were compared with untreated cells and statistically evaluated using the Student's *t*-test. (**A**) RhoA, (**B**) RhoB. (\* *p* < 0.05; \*\* *p* < 0.01; \*\*\* *p* < 0.001, *n* = 3).

**Table 1.** Summary of the results.

\* 1: PD-L2 mRNA was suppressed and increased by LMF in HT-1080 cells and TIG-1 cells, respectively. \* 2: PC-9 cell mRNA level without LMF was set to 1.00 and the relative expression levels for other mRNAs were calculated. **–**: not tested.

### **3. Discussion**

In the present study, we examined the effects of LMF on six gene products including PD-L1, PD-L2, EGFR, VEGF, RhoA, and RhoB, involved in immune evasion, angiogenesis, and malignant transformation, using the human fibrosarcoma HT1080 cell line. We reveal that LMF downregulated five gene products, while upregulating RhoB expression.

One of the important determinants to improve the efficacy of cancer therapy is inhibiting cancer cell evasion from the host tumor immune surveillance system [8]. Various tumors express increased levels of PD-L1 to evade such host defense mechanisms and eventually facilitate tumor growth [2,4,8,9]. Moreover, the tumor growth in terms of Breslow tumor thickness in melanoma specimens correlated with the level of PD-L1 expression, and the survival rate of patients expressing a high-level of PD-L1 was statistically lower than that of low-expressing patients with stage II melanoma [5]. Therefore,

we confirmed the effects of LMF on the downregulation of PD-L1 expression in cancer cells. We first examined PD-L1 mRNA levels in six cancer cell lines, HT1080, MCF-7, PC-9, NIH:OVCAR-3, PANC-1, and A549 cells, and in one normal cell line TIG-1. All cell lines express PD-L1 levels similar to or greater than that of the A549 cell line, a low-expressing control [12,58]. The PD-L1 mRNA levels expressed by TIG-1 cells were comparable to those of the high-expressing control cell line PC-9 [12,44,54]. This observation agrees with earlier reports on several human cells, including lung cells and various tissues, which express low- to high-levels of PD-L1 [3,59,60]. These data strongly suggest the importance of selective inhibition of PD-L1 expressed on cancer cells but not on normal cells.

Among the cell lines tested, the HT1080 cell line expressed the highest levels of PD-L1 mRNA and was a suitable model to test the suppressive effects of LMF with emphasis on PD-L1 expression at the mRNA and protein levels. Several inherent characteristics of the HT1080 cell line are known since its establishment in 1974 [61]. HT1080 cells contain both wild-type and mutant N-Ras alleles and are expressed at the protein level. The mutant N-Ras protein induces cell transformation in cells even though it is present in far less amounts compared to that of normal p21 N-Ras in parental HT1080 cells [62]. The mutant N-Ras protein maintains a chronically activated state due to constitutive GTP binding, thereby conferring the constitutive activation of Ras-dependent signaling pathways. Representative Ras-regulated pathways are the Raf, Rac1, RhoA, and PI3K signal cascades, which are known to regulate mitogenesis, motility and invasiveness, actin cytoskeletal architecture, and cell survival, respectively [19,63]. Among the intermediary transducers, activated Raf/MEK is necessary for the aggressive tumorigenic phenotype in HT1080 cells but not the PI3K/Akt pathway [18,19]. However, others reported that the activated mutant N-Ras protein constitutively stimulates the Raf/MEK and the PI3K/Akt pathways to induce PD-L1 gene expression in several cancer cell lines [6,17].

Although, these data suggest that PD-L1 expression could be driven by the Ras regulated pathways in HT1080 cells, direct data proving an association between the Ras regulated pathways and PD-L1 expression in HT1080 cells are not available to date. In the present study, we show significant suppression of mRNA, protein and surface protein levels of PD-L1 in HT1080 cells by LMF. These results show suppression of activated mutant N-Ras regulated pathways for PD-L1 expression and existing data show many transducers in the Ras regulated pathways being suppressed by fucoidan [20–22,39]. LMF down-regulation of PD-L1 expression via Ras pathways in HT1080 cells could be explained by the possible regulatory pathways proposed for PD-L1 gene expression in cancer cells. According to this model, growth factor (GF)-stimulated pathways such as Ras/Raf and Ras/PI3K drive PD-L1 expression [6]. Thus, these pathways for PD-L1 expression coincide mostly with the Ras-regulated pathways proposed in HT1080 cells, as above. Although, the mutant N-Ras protein is constitutively active in regulating the pathways involving Raf, PI3K, and Rac transducers, the normal N-Ras-regulated pathways activated by GFs should be evaluated separately because HT1080 cells express functional wild-type and mutant N-Ras proteins [62].

Normal N-Ras protein is activated by exogenous stimuli such as by the binding of EGF, VEGF, and platelet-derived growth factor (PDGF) with their receptors (EGFR, VEGFR, and PDGFR). These transmembrane receptors mediate GTP-binding with the normal N-Ras protein for activation, enabling regulation of downstream transducers [63]. Cellular activities including proliferation, growth, survival, angiogenesis, metastases, and motility are regulated by the activated EGFR, VEGFR, and PDGFR through pathways such as the mitogen-activated protein kinase (MAPK) pathway: Ras/Raf, the PI3K pathway: PI3K/Akt, and the JAK/STAT pathway: JAK/STAT [64]. Available data shows that EGF-stimulated EGFR upregulates PD-L1 expression through EGFR/PI3K/Akt, EGFR/Ras/Raf, and EGR/PLC-γ signaling pathways [2,12,65,66]. Moreover, tumor cells can evade the immune system or promote their own growth through increased EGFR expression and indoleamine 2,3-dioxygenase 1 or VEGF production [67]. Although ligand-bound-activated EGFR with increased protein expression was present in HT1080 cells [14,68], the supernatant collected from the HT1080 cell culture did not contain secreted EGF but contained moderate concentrations of TGF-α which correlated with pEGFR expression [14]. Thus, an autocrine loop of TGF-α activates wild-type EGFR as well as the downstream

factors in HT1080 cells, as both EGF and TGF-α belong to eight members of EGF family ligands, and TGF-α is an EGFR ligand in cancer [69,70]. Similarly, it has been reported that anti-EGF receptor mAbs prevented EGF and TGF-α induced growth stimulation in many human cells [14,71]. Accumulated data suggest that inhibition of EGFR or VEGF could inhibit HT1080 cell growth. In this study, we report significant suppression by LMF of both EGFR transcription and cell growth, suggesting a mechanism to suppress PD-L1 expression through regulation of EGFR expression. A plausible explanation for EGFR suppression by LMF would be that fucoidan binds with EGF causing prevention of EGF-induced phosphorylation of EGFR [72,73]. In support of this, earlier reports showed that fucoidan from *Laminaria guryanovae* exerts a potent inhibitory effect on EGF-induced phosphorylation of EGFR in mouse epidermal JB6 Cl41 cells [72] and fucoidan KW derived from brown algae *Kjellmaniella crassifolia* caused significant reduction in the levels of phosphorylated EGFR and PKCα proteins in A549 cells [43]. Others have shown that fucoidan suppresses growth factor-stimulated pathways involving the Ras/Raf pathway and inhibits the phosphorylation of intermediary transducers such as ERK1/2, PI3K, Akt, mTOR, c-Jun, c-Fos, EGFR, and VEGF [21,22,39]. While future studies are needed to confirm whether LMF binds with TGF-α in reducing EGFR phosphorylation, it is probable to suggest that reduction in the EGFR mRNA levels by LMF decreases the translated and phosphorylated EGFR levels. Contrary to others and ours, an existing report shows that EGFR could not be detected in HT1080 cells and suggested that HT1080 cell oncogenic signaling through Ras/PI3K/Akt is independent of EGFR [74]. In addition, fucoidan exerted a potent inhibitory effect on EGF-induced phosphorylation of EGFR and phosphorylation of ERK and JNK under the control of EGF. Furthermore, EGF-induced c-*fos* and c-*jun* transcriptional activities were inhibited by fucoidan leading to inhibition of AP-1 activity and cell transformation induced by EGF [72].

VEGF is present in a number of cancer cell lines including HT1080 cells and is suggested to play a crucial role in HT1080 cell metastasis based on results obtained using the anti-human VEGF antibody, which reduced spontaneous lung micrometastasis [75]. Moreover, inhibition of tumor-secreted VEGF limits primary tumor growth of sarcoma cell lines by inhibiting host angiogenesis [76]. An inhibitory mechanism of angiogenesis by low molecular weight fucoidan (LMWF), which is distinct from LMF, inhibits hypoxia-induced reactive oxygen species (ROS) formation, HIF-1α expression, VEGF secretion, and downstream VEGFR2/PI3K/Akt/mTOR/p70S6K1/4EBP-1 cascade in human bladder T24 cancer cells, ultimately suppressing HIF-1α/VEGF transcription and angiogenesis [39]. VEGF upregulates both PD-L1 mRNA and protein expression [2,77]. Thus, suppression of both VEGF-enhanced angiogenesis and PD-L1 expression contributes to inhibiting cancer cell evasion. Supporting evidence shows that fucoidan extract inhibits expression and secretion of VEGF which suppresses vascular tubule formation in HeLa cells and suppressing invasion of HT1080 cells [41]. In the present study, because LMF caused significant suppression of both VEGF mRNA and PD-L1 mRNA/protein levels in HT1080 cells, we can extrapolate that cancer cells become more susceptible to the host defense system. The rationale behind the suppression of the PD-L1 level by fucoidan comes from the fact that fucoidan directly binds with VEGF, thereby preventing VEGFR phosphorylation [70,78]. Other growth factors, PDGF and PDGFR, are constitutively expressed in HT1080 cells. PDGF-A is constitutively secreted into the medium, where it binds to and activates PDGFR and downstream PI3K and Akt pathways [19]. Although we did not test the suppressive effect of LMF on PDGF expression, there are studies showing that fucoidan suppresses PDGF expression. Oligo-fucoidan suppresses PDGF-induced cell proliferation and induces G1/G0 cell cycle arrest in airway smooth muscle cells [79]. Therefore, LMF is likely to aid in HT1080 cell growth inhibition by suppressing PDGF expression. As mentioned thus far, LMF suppresses VEGF and EGFR expression in HT1080 cells to suppress Ras-regulated pathways for PD-L1/PD-L2 expression.

The Ras superfamily contains a subgroup of Rho GTPases composed of 20 kinds of intracellular signaling molecules including RhoA and RhoB proteins. Constitutively expressed RhoA functions in tumor development. Indeed, RhoA and its downstream pathway is constitutively active in HT1080 cells. The present study shows significant suppression of RhoA mRNA levels by LMF, suggesting that LMF downregulates Ras-controlled oncogenic activities via the RhoA pathway [19]. In contrast, the role of RhoB is not clearly known, it acts as either an oncogene under some conditions or a tumor suppressor gene under others [80,81]. RhoB expression is induced by EGF, PDGF, and many other agents, while several factors including N-Ras and EGFR suppress RhoB promoter activity in NIH3T3 cells and human cancer cell lines derived from lung, pancreatic, and cervical tumors [82]. Fucoidan suppresses PDGF (RhoB inducer) expression indicating downregulation of RhoB expression by fucoidan [79]. Conversely, LMF suppresses EGFR (RhoB suppressor), and fucoidan suppresses phosphorylation of various Ras-regulated transducers. Together these data suggest that LMF/fucoidan could act as an upregulator of RhoB because LMF significantly and dose-dependently upregulated RhoB mRNA levels, supporting the role of RhoB as a tumor suppressor. Thus, LMF performs a significant role in antitumor effect by upregulating RhoB expression in several cancer types [80,81]. Suppressive effects by fucoidan on Ras-regulated pathways are presented in Figure 11.

**Figure 11.** Low molecular weight fucoidan extract (LMF) and several fucoidan species regulate multiple transcription factors in the Ras-controlled pathways in various cells, including HT1080 cells. The numbers in the square bracket indicate the reference numbers, and downward and upward arrows indicate the factors up- and down-regulated by various fucoidan species reported in the references and LMF used in this study. Adapted from [2,6,19].

An important finding that LMF neither inhibited PD-L1 expression nor growth of the TIG-1 cells suggests selective activity towards cancer cells expressing PD-L1. Similar to our results with normal cells, another report shows a lack of effect of fucoidan on normal human mammary epithelial cell growth [83]. These results collectively suggest that LMF/fucoidan treatment against PD-L1 expression is effective because of its selective action on cancer cells. However, the anti-PD-L1 antibody treatment may not be efficacious because not all cancer patients express higher PD-L1 [67]. Similarly, the suppressive effect on PD-L1 expression conferred by LMF/fucoidan may not be efficacious for patients who are not expressing higher PD-L1 [67]. Thus, as both methods rely on the expressed PD-L1 or its expression in cancer cells, these methods have a common limitation in using PD-L1 as a treatment target. On the other hand, in addition to PD-L1 suppression by LMF, extensive data exists demonstrating that fucoidan exhibits various biological activities including the suppressive modulation of the intracellular transducers that regulate proliferation, growth, survival, angiogenesis, metastases, and motility in

cancer cells [22,36,39,42,43,84]. Therefore, LMF/fucoidan exhibits anticancer effects due to the broad biological activities independent of its anti-PD-L1 effect.

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

### *4.1. Cell Lines and Cell Culture*

Seven established human cell lines and their culture conditions are shown in Table 2.

**Table 2.** Human cell lines and culture conditions.

MEM (Eagle's Minimum Essential Medium, Nissui Pharmaceutical Co., Ltd., Tokyo, Japan). HEPES (Dojindo Laboratories, Kumamoto, Japan), NEAA (Non-Essential Amino Acids, FUJIFILM Wako Pure Chemical Corporation, Osaka, Japan). Penicillin G Potassium (Meiji Seika Pharma Co., Ltd., Tokyo, Japan). Streptomycin sulfate (Meiji Seika Pharma). FBS: (fetal bovine serum, HyClone, Thermo Fisher Scientific Inc., Waltham, MA, USA). DMEM (Dulbecco's Modified Eagle's Medium, Nissui Pharmaceutical, Tokyo, Japan). RPMI (Nissui Pharmaceutical, Tokyo, Japan). #1 ATCC: American Type Culture Collection, Manassas, VA, USA; #2 IDAC: The Cell Resource Center for Biomedical Research, Institute of Development, Aging and Cancer, Tohoku University, Sendai, Japan; #3 RIKEN: BioResource Research Center, Ibaraki, Japan.

### *4.2. Fucoidan*

The abalone glycosidase-digested LMF, commercially available as "Power fucoidan," was generously donated for this study by the Daiichi Sangyo Corporation (Osaka, Japan). LMF was prepared as previously described [2]. Briefly, high molecular weight fucoidan extract from seaweed of *Cladosiphon novae-caledoniae* Kylin was purified to 85% purity and digested with glycosidases to obtain LMF. LMF consists of a digested small molecular weight fraction (72%, MW < 500 Da) and non-digested fractions (less than 28%, peak MW: 800 kDa). LMF consisted mostly of fucose (73%), xylose (12%), and mannose (7%). The ratio of sulfation was 14.5%.

### *4.3. WST-1 Assay (Cell Viability Assay)*

WST-1 assay is commonly used to measure the number of live cells after treating the cells with culture containing test agents [53]. This assay was used to measure the cytocidal effects of LMF on HT1080 cells. 100 <sup>μ</sup>L each of HT1080 (2 <sup>×</sup> <sup>10</sup><sup>4</sup> cells/mL) and TIG-1 (4 <sup>×</sup> <sup>10</sup><sup>4</sup> cells/mL) cells were plated in 96-well microplates and cultured for 24 h at 37 ◦C. The WST-1 measurement was performed according to the standard protocol described by the manufacturer. Briefly, the cells were exposed to various concentrations of LMF for 48 h. Spent medium was removed and cells were washed once with PBS. To each well, 100 μL of culture medium containing 0.5 mM WST-1 (Dojindo) and 0.02 mM 1-methoxy phenazine methosulfate (Dojindo) was added and incubated for 1 h at 37 ◦C. Cell metabolic activity was measured at 450 nm using a microplate reader (Tecan, Männedorf, Switzerland).

### *4.4. Hoechst Staining (Cellular DNA Staining for Cell Number Determination)*

Hoechst 33342 dye stains both live and apoptotic cell DNA [50,51]. It is possible to estimate the cytocidal effect of LMF by analyzing WST-1 and Hoechst Staining assays. The cells in the same microplates used for WST-1 assay were fixed with 4% formaldehyde solution (Wako) for 30 min at 25 ◦C. Cells were washed once with PBS and stained with 100 μL PBS per well, containing 2 μg/mL of Hoechst 33342 (Dojindo), for 30 min in the dark at 25 ◦C, according to the instruction manual. Fluorescence intensity (excitation, 360 nm and emission, 465 nm) was measured by the multimode plate reader, infinite F200 PRO (Tecan).

### *4.5. Immuno-Fluorescence Staining*

Previous studies have used the Immuno-fluorescence staining method to see the effect of fucoidan on T24 cells [39]. The same method was also used to detect PD-L1 in the cultured cells [58]. Therefore, we used this method to detect PD-L1 expressed in seven cell lines. HT1080 and A549 cells at <sup>2</sup> <sup>×</sup> <sup>10</sup><sup>4</sup> cells/mL, MCF-7, PC-9, NIH:OVCAR-3, and PANC-1 cells at 3 <sup>×</sup> <sup>10</sup><sup>4</sup> cells/mL, and TIG-1 cells at <sup>4</sup> <sup>×</sup> <sup>10</sup><sup>4</sup> cells/mL were prepared, and 100 <sup>μ</sup>L of each cell line was seeded per well in a 96-well black plate (Greiner Bio-One International GmbH, Kremsmünster, Oberösterreich, Austria) and cultured for 24 h. Cells were fixed with 4% paraformaldehyde (Wako) for 15 min. After washing with PBS, cells were treated with blocking solution containing 0.1% Tween-20 with 1% BSA in PBS for 1 h. After washing three times with PBS, cells were treated overnight at 4 ◦C with the anti-PD-L1 Rabbit mAb (E1L3N, Cell Signaling Technology Co., Danvers, MA, USA) diluted 800-fold with blocking solution. Rabbit mAb IgG Isotype Control (DA1E, Cell Signaling Technology) was diluted 800-fold with blocking solution and used as an isotype antibody control. After washing three times with PBS, cells were treated for 30 min with the secondary antibody, anti-Rabbit IgG (H+L), F(ab')2 Fragment (Alexa Fluor® 555 Conjugate) (Cell Signaling Technology), and diluted 1,000-fold with blocking solution. Cells were washed one time with PBS and stained with 2 μg/mL of Hoechst 33342 for 30 min. Fluorescence intensity of each cell line was measured using IN Cell Analyzer 1000 (GE Healthcare UK Ltd., Buckinghamshire, UK).

### *4.6. Quantitative Reverse Transcription Polymerase Chain Reaction (qRT-PCR)*

Polymerase chain reaction (PCR) has been used broadly, to detect a specific gene transcript [85], and to quantitate the target transcript level [86,87]. Quantitative detection and analyses of the expressed gene transcripts have been advanced greatly by incorporating automated instrumentations. Seven gene transcripts were examined by fully utilizing the commercially available instruments and reagents, as described below. Five milliliters of HT1080 cells were seeded at a density of 5 <sup>×</sup> 10<sup>4</sup> cells/mL per 60 mm dish (FALCON, Corning Inc., Corning, NY, USA) and cultured for 24 h. Spent medium was replaced with the medium containing various LMF concentrations (0, 1, 10, or 100 μg/mL). After 48 h of treatment, total RNA was isolated using a High Pure RNA Isolation Kit (F. Hoffmann-La Roche Ltd., Basel, Switzerland) following the manufacturer's protocol. cDNA was synthesized using total RNA and ReverTra Ace qPCR RT Kit Master Mix with gDNA Remover (TOYOBO Co., Ltd., Osaka, Japan) and was used for RT-PCR templates. For specific gene transcript amplification, cDNA template, primer pairs (Table 3), and THUNDERBIRDTMSYBR® qPCR Mix (TOYOBO) mixture were placed in a 96-well PCR plate (NIPPON Genetics Co., Ltd., Tokyo, Japan) in triplicate. Primer sequences for amplifying the transcripts of seven genes (EGFR, VEGF, RhoA, RhoB, PD-L1, PD-L2, and GAPDH), shown in Table 1, were purchased from Sigma-Aldrich Co. (St. Louis, MO, USA). Quantitative reverse transcription polymerase chain reaction (qRT-PCR) for seven genes was carried out using Thermal Cycler DiceTM Real Time System TP800 (Takara Bio Inc., Shiga, Japan). The thermal program was as follows: denaturation at 95 ◦C for 30 s, 45 cycles of denaturation at 95 ◦C for 5 s, annealing at 60 ◦C

for 10 s, and extension at 72 ◦C for 20 s. Fluorescence values for each gene were normalized using GAPDH as an internal standard and used for statistical evaluation.

**Table 3.** Primer sequences used for qRT-PCR analyses.

### *4.7. Flow Cytometry (FCM) Analysis*

Five milliliters of 3.1 <sup>×</sup> 103 cells/mL of HT1080 cells were plated on 60 mm dish and incubated for 24 h. The cells were then treated with 5 mL of medium with or without 10 μg/mL of LMF for 5 days. Cells were dislodged from the dish by treating with 0.05% Trypsin-0.02% EDTA solution (Thermo Fisher Scientific) for 3 min at 37 ◦C. Cells were fixed with 4% paraformaldehyde for 10 min at 37 ◦C, collected, and treated for 1 h with the drawback Rabbit mAb (E1L3N) or with the corresponding Rabbit mAb IgG isotype control (DA1E) diluted 800-fold with incubation buffer. After washing three times with PBS, cells were treated for 30 min with the secondary antibody, anti-Rabbit IgG (H+L), F(ab')2 Fragment (Alexa Fluor® 488 Conjugate) (Cell Signaling Technology) diluted 500-fold with incubation buffer. Fluorescence intensity of each cell line was measured using Flow Cytometer EPICS XL System II-JK (Beckman Coulter, Inc., Brea, CA, USA) which integrates flow cytometry technology, thus enabling the study of cellular populations [92]. Data analyses were carried out using Flow Cytometry (FCM) analysis software, FlowJo (Version 4.6.2) (Tree Star, Inc., Ashland, OR, USA) and the results expressed as histograms and MFI.

### *4.8. Western Blot Analysis*

In our previous study, the Western Blot method was used to analyze the expression of Bcl-2 family proteins [47]. We used this method to detect PD-L1 in HT1080 cell lysates treated with LMF. HT1080 cells were cultured for 5 days in the medium supplemented with or without 10 μg/mL of LMF. Cells were washed three times with cold PBS and lysed for 5 min with 400 μL of protein extraction buffer (M-PER Mammalian Protein Extraction Reagent (Thermo Fisher Scientific) supplemented with 1% of Halt Protease Inhibitor Single-Use Cocktail (Thermo Fisher Scientific) and 5 mM EDTA). Lysates were collected in 1.5 mL microtubes and cell lysis was further ensured by repeated sonication for four times at 5 s intervals using the Handy Sonic model UR-20P (TOMY SEIKO CO., LTD., Tokyo, Japan). Protein extracts were centrifuged at 14,000× *g* for 15 min at 4 ◦C. Supernatants were recovered and stored at −80 ◦C. Prior to use, the protein concentration was determined using the Bradford protein assay. Supernatants were diluted 50-, 100-, 200-, and 400-fold with PBS. A standard solution was prepared using bovine γ globulin (1 mg/mL) by serially diluting with PBS from 90 to 0 μg/mL with 10 μg/mL decrement. To determine the protein concentrations in the dilution series, 100 μL each of the protein

extracts and standard solutions were mixed with 100 μL of 2.5-fold diluted Bio-Rad protein assay dye reagent concentrate (Bio-Rad Laboratories, Inc., Hercules, CA, USA) in a 96-well microplate (Nunc, Thermo Fisher Scientific), and absorbance at 595 nm were measured using a microplate reader (TECAN). Protein concentrations in the treated HT1080 cell extracts were determined based on the standard curve derived from bovine γ globulin. Equal amounts of protein from each sample were separated by electrophoresis through SDS-PAGE and transferred to Amersham Hybond P polyvinylidene fluoride 0.45 membranes (GE Healthcare) using The Trans-Blot® SD semi-dry transfer cell apparatus (Bio-Rad) following the manufacturer's protocol. Membranes were blocked for 1 h at room temperature with gentle shaking in Tris-buffered saline containing 5% skim milk powder (Wako). Membrane was cut into two pieces so as to apply different antibodies for detecting closer molecular weight proteins of GAPDH (37 kD) and PD-L1 (40-50 kD) unambiguously. The membrane was incubated overnight at 4 ◦C with the primary antibody, anti-PD-L1 Rabbit mAb (E1L3N) (Cell Signaling Technology) or anti-GAPDH Rabbit mAb (D16H11) (Cell Signaling Technology), diluted 1000-fold or 4000-fold respectively, in Solution 1 of Can Get Signal® Immunoreaction Enhancer Solution (Toyobo). The membranes were washed twice and incubated for 1 h at room temperature with the Anti-Rabbit IgG, horseradish peroxidase (HRP)-linked secondary antibody (Cell Signaling Technology), diluted 2000-fold with Solution 2 of Can Get Signal® Immunoreaction Enhancer Solution. Chemiluminescence signals of protein bands obtained by ECL prime western blotting detection reagent (GE healthcare), as described by the supplier, were detected with LAS-1000 (FUJIFILM Corporation, Tokyo, Japan), which were quantified and numerated using image J (1.4.6r) software.

### *4.9. Inhibitory Activity of LMF on PD-1:PD-L1 Binding*

Inhibitory activity of LMF on PD-1 and PD-L1 binding was examined using PD-1:PD-L1(Biotinylated) Inhibitor Screening Colorimetric Assay Kit (BPS Bioscience, San Diego, CA, USA) following the manufacturer's protocol. Briefly, PD-1 protein was coated on a 96-well plate and added biotinylated PD-L1 to form complex of PD-1:PD-L1:biotin. The plate was treated with Streptavidin-HRP followed by the addition of a colorimetric HRP substrate to produce color. Thus, this set up will serve as a positive control. To this set up, the addition of PD-1 neutralizing antibody as a test inhibitor will reduce color production serving as a second positive control. When a prospective inhibitor such as LMF is added to this set up, color production is reduced only when LMF binds to PD-1. For ligand control, the well is not coated with PD-1 and for the blank control well, biotinylated PD-L1 is not included. After incubation, these set-ups produce appropriate colors, which can be measured at 450 nm using a microplate reader (TECAN).

### *4.10. Statistical Analysis*

Each experiment was performed at least in triplicate and repeated three times. The results are presented as the mean ± standard deviation (SD) values. The difference between the two groups was analyzed using the two-tailed Student's *t*-test, and differences among three or more groups were analyzed using one-way analysis of variance (ANOVA) with Tukey's multiple comparisons. Statistical analysis was done using Mini StatMate (ATMS Co., Ltd., Tokyo, Japan). A value of *p* < 0.05 represents a significant difference (\* *p* < 0.05; \*\* *p* < 0.01; \*\*\* *p* < 0.001).

### **5. Conclusions**

Our findings revealed that LMF exerts its effect by specifically suppressing not only mRNA levels but also cytoplasmic- and surface-protein levels of PD-L1 in HT1080 fibrosarcoma cells. We also provide data for the downregulation of VEGF, EGFR, and RhoA mRNAs, and the upregulation of RhoB mRNA in HT1080 cells. Together, these data suggest that LMF could be used as a complementary agent in the treatment of various types of cancers that express high PD-L1 levels.

**Author Contributions:** K.T. conceived and designed the experiments; Y.K. and K.T. performed the experiments and analyzed the data; H.E. and N.N. supplied reagents/materials; K.T. and S.S. analyzed the data and wrote the manuscript.

**Funding:** This research was funded by JSPS KAKENHI, Grant Number 17K07793, and was partially supported by Daiichi Sangyo Corporation (Osaka, Japan).

**Conflicts of Interest:** This research was partially supported by Daiichi Sangyo Corporation. The funders had no role in the design of the study, in the collection, analyses, or interpretation of data, in the writing of the manuscript, or in the decision to publish the results.

### **References**

© 2019 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 (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **Actinomycin V Inhibits Migration and Invasion via Suppressing Snail**/**Slug-Mediated Epithelial-Mesenchymal Transition Progression in Human Breast Cancer MDA-MB-231 Cells In Vitro**

### **Shiqi Lin 1, Caiyun Zhang 1, Fangyuan Liu 1, Jiahui Ma 1, Fujuan Jia 1, Zhuo Han 1, Weidong Xie <sup>1</sup> and Xia Li 1,2,3,\***

Received: 23 April 2019; Accepted: 22 May 2019; Published: 24 May 2019

**Abstract:** Actinomycin V, an analog of actinomycin D produced by the marine-derived actinomycete *Streptomyces* sp., possessing a 4-ketoproline instead of a 4-proline in actinomycin D. In this study, the involvement of snail/slug-mediated epithelial-mesenchymal transition (EMT) in the anti-migration and -invasion actions of actinomycin V was investigated in human breast cancer MDA-MB-231 cells in vitro. Cell proliferation effect was evaluated by 3-(4,5-Dimethylthiazol)-2,5-diphenyltetrazolium bromide (MTT) assay. Wound-healing and Transwell assay were performed to investigate the anti-migration and -invasion effects of actinomycin V. Western blotting was used to detect the expression levels of E-cadherin, N-cadherin, vimentin, snail, slug, zinc finger E-box binding homeobox 1 (ZEB1), and twist proteins and the mRNA levels were detected by rt-PCR. Actinomycin V showed stronger cytotoxic activity than that of actinomycin D. Actinomycin V up-regulated both of the protein and mRNA expression levels of E-cadherin and down-regulated that of N-cadherin and vimentin in the same cells. In this connection, actinomycin V decreased the snail and slug protein expression, and consequently inhibited cells EMT procession. Our results suggest that actinomycin V inhibits EMT-mediated migration and invasion via decreasing snail and slug expression, which exhibits therapeutic potential for the treatment of breast cancer and further toxicity investigation in vivo is needed.

**Keywords:** breast cancer; actinomycin; EMT; migration; invasion

### **1. Introduction**

Breast cancer has the highest diagnosis and death rates among women followed by lung and colorectal cancer and has generally increased in recent years [1]. Since many therapies including surgery, radiotherapy, chemotherapy, and human epidermal growth factor receptor 2 (HER2) targeted therapy have been used in clinical trials, the overall survival rate of patients still remained poor [2]. Metastasis is the majority reason for breast cancer death. Epithelial-mesenchymal transition (EMT) is a key process in cancer invasion and metastasis [3]. EMT is a dynamic and complex multistep process that cells attach to and degrade the surrounding extracellular matrix (ECM) to escape from the primary tumor site, invade the stromal tissues, and establish the distant secondary tumor [4]. Therefore, attention has been focused on blocking the migration and invasion processes of breast cancer cells to inhibit metastasis [5–7].

Many effective anti-cancer compounds that were originally isolated from marine invertebrates, such as discodermolide, halichondrin B, bryostatin1, and phorboxazole A, were actually produced by marine microorganisms [8–10]. Marine microorganisms were considered to be the promising sources for discovering new effective anti-cancer drugs and many efforts have been made in recent years. Actinomycins, produced by the marine-derived actinomycete *Streptomyces* sp., is a class of cyclic chromopeptide consisting of two cyclic pentapeptide lactones attached to a central phenoxazinone chromophore via amide bonds [11–13]. Typically, actinomycin D has been well used in treatment of a variety of cancers such as Wilms tumor, rhabdomycosarcoma, and Ewing's sarcoma through binding and intercalating double-stranded DNA and inhibiting RNA synthesis [14]. A report has shown that actinomycin D inhibited the migration of MDA-MB-231 cells but the underlying mechanism was not mention [15]. Moreover, the toxicity development, particularly in hepatotoxicity of actinomycin D, is associated with use in an anti-tumor process, which severely limits its effectiveness in clinic. Actinomycin V possesses 4-ketoproline as a substitute for 4-proline of actinomycin D. Studies have reported that actinomycin V's inhibitory effects on the F5-5 friend leukemia cells, MCF-7, A549, and K562 cells were superior to actinomycin D and the effect on human breast cancer cells MCF-7 was the most obvious [16,17]. However, the anti- migration and invasion actions of actinomycin V and the underlying mechanism remains unclear. In this present study, we examined that actinomycin V treatment inhibited the proliferation of human breast cancer cells and suppressed the EMT process by down-regulating the snail and slug protein expression. These results may provide a new theoretical basis for the treatment of breast cancer with actinomycin V and the useful information for developing novel marine-derived anti-cancer drugs.

### **2. Results**

### *2.1. Actinomycin V Inhibits the Proliferation of Human Breast Cancer Cells*

To compare the effects of three actinomycins (actinomycin D, actinomycin V, and actinomycin Xob as shown in Figure 1) on breast cancer cells, we initially performed 3-(4,5-Dimethylthiazol)-2,5-diphenyltetrazolium bromide (MTT) analysis in several subtypes of human breast cancer (MDA-MB-231, BT474, and MCF-7) and noncancer (HMLE and MCF-10A) cell lines. We found that actinomycin V had the excellent activity on breast cancer cells among three actinomycins. Actinomycin V displayed the strongest sensitivity on MDA-MB-231 cells and with lower cytotoxicity in noncancer breast cells as shown in the Table 1. The IC50 of actinomycin V for 48 h treatment to MDA-MB-231, MCF-7, BT474, HMLE, and MCF-10A were 0.83 ± 0.32 nmol/L, 1.92 ± 0.25 nmol/L, 4.16 ± 0.25 nmol/L, 3.49 ± 0.31 nmol/L, and 4.07 ± 0.26 nmol/L, respectively, and actinomycin D treatment for 48 h to those cell lines were 15.15 ± 0.52 nmol/L, 8.23 ± 0.50 nmol/L, 37.00 ± 3.15 nmol/L, 30.22 ± 0.50 nmol/L, and 34.01 ± 0.25 nmol/L, respectively. Further experiments showed that actinomycin V dose- and time-dependently inhibited the proliferation of human breast cancer cells (Figure 2).

**Figure 1.** Chemical structure of actinomycin D, actinomycin V, and actinomycin Xob.

**Table 1.** Growth inhibitory activities of three actinomycins and Adriamycin in different cell lines.

<sup>1</sup> The effects of three actinomycins and Adriamycin on various human breast cancer cells (MCF-7, MDA-MB-231, and BT474) and normal human breast epithelial cell lines (HMLE and MCF-10A) were examined by 3-(4,5-Dimethylthiazol)-2,5-diphenyltetrazolium bromide (MTT) assay. The cells were treated with various concentrations of compounds for 48 h and the IC50 values were then calculated. Results were obtained from three independent experiments. Data were expressed as the mean ± SD (Standard Deviation) of triplicate experiments.

**Figure 2.** Actinomycin V does- and time- dependently inhibited the proliferation of various cell lines. Cells were treated with various concentrations of actinomycin V (0 to 5 nmol/L) for 24 h to 72 h. Cell viability was denoted as a percentage of the untreated control (actinomycin V 0 nmol/L) at the concurrent time point. Results were obtained from three independent experiments.

### *2.2. Actinomycin V's E*ff*ects on Morphological Changes in Breast Cancer Cells*

We observed the effects of actinomycin V on morphological changes in breast cancer cells (MCF-7, MDA-MB-231, and BT474) (Figure 3). After 1 nmol/L actinomycin V 24 h treatment cells performed in different levels of morphological changes. Especially the metastatic MDA-MB-231 cells, it displayed in a scattered distribution in the culture and a spindle or star-like structure before actinomycin V treated but after 24 h treatment cells changed to a round cell shape and tend to an aggregation distribution, indicating cells appeared to have improved intercellular adhesion and reduced cells migration and invasion.

**Figure 3.** Changes in morphology of MCF-7, MDA-MB-231, and BT474 cells upon actinomycin V treatment. (**A**–**C**) Cells were treated with 1 nmol/L actinomycin V for 24 h and phase contrast images of the cultured cells were taken under 200× objective lenses.

### *2.3. Actinomycin V Inhibits the Migration and Invasion of Human Breast Cancer Cells*

Considering that migration and invasion are the major drivers of breast cancer metastasis, wound-healing assay and Transwell assay were used to further confirm the actinomycin V effects on anti-migration and anti-invasion in human breast cancer cells. In the wound-healing assay (Figure 4), the untreated MDA-MB-231 cells wound-healing rate was 52% in 24 h. However, actinomycin V treatment apparently retarded the wound healing, the healing rates were only about 23% and 10%, respectively, in the presence of 1–2 nmol/L actinomycin V. In the Transwell assay (Figure 5), actinomycin V dose-dependent treatment significantly suppressed the invasion of MDA-MB-231 cells. Similarly, actinomycin V treatment inhibited the migration and invasion of MCF-7 and BT474 cells but the inhibitory effects were not as significant when compared with its anti-migration and anti-invasion effects on the metastatic MDA-MB-231 cells. These results indicated that actinomycin V inhibited the migration and invasion of human breast cancer cells.

**Figure 4.** Effect of actinomycin V on the motility of MDA-MB-231, MCF-7, and BT474 cells. The wound-healing rate of each cell line was analyzed (magnification: ×200). Results were obtained from three independent experiments. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001.

**Figure 5.** Effects of actinomycin V on the invasion of MDA-MB-231, MCF-7, and BT474 cells. (**A**,**B**) Cells were treated with 0–2 nmol/L actinomycin V for 24 h, and the invasion in these two cell lines was analyzed (magnification: ×200). Results were obtained from three independent experiments. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001.

### *2.4. E*ff*ects of Actinomycin V on the Expression of EMT-Associated Proteins and mRNA in MDA-MB-231 Cells*

To investigate the actinomycin V effects on Epithelial-mesenchymal transition inhibition in human breast cancer cells, we preformed the Western blot analysis to evaluate the expression levels of the epithelial marker E-cadherin, and mesenchymal markers N-cadherin and Vimentin in MDA-MB-231 cells (Figures 6 and 7). We found that actinomycin V treatment increased the expression levels of E-cadherin while decreased that of N-cadherin and Vimentin. Immunofluorescence staining also showed that compared with untreated cells, actinomycin V treatment significantly decreased intracellular N-cadherin and vimentin content while increased the E-cadherin content in cancer cells.

**Figure 6.** Effect of actinomycin V on the expression of E-cadherin. (**A**,**B**) MDA-MB-231 cells were treated with 0–2 nmol/L actinomycin V for 24 h then the protein expression of E-cadherin was measured by Western blot. (**C**) Relative expression of E-cadherin mRNA in the MDA-MB-231 cells was analyzed by real-time PCR. RNA levels are represented as fold increase relative to the level of the control (normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA level). Results were obtained from three independent experiments. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001. (**D**) cells were treated with 0–2 nmol/L actinomycin V for 6 h and analyzed by E-cadherin fluorescent signals. Cells were stained with anti- E-cadherin (green) and 4 ,6-diamidino-2-phenylindole (DAPI, blue). Magnification ×200. E-cadherin levels were increased in the cells, consistent with the Western blot results.

**Figure 7.** Effects of actinomycin V on the N-cadherin and vimentin expression. (**A**–**C**) MDA-MB-231 cells were treated with 0–2 nmol/L actinomycin V for 24 h then the protein expression of vimentin and N-cadherin was measured by Western blot. (**D**) cells were treated with 0–2 nmol/L actinomycin V for 6 h and analyzed by vimentin and N-cadherin fluorescent signals. Cells were stained with anti-vimentin (red), N-cadherin (green), and DAPI (blue). Magnification: ×200. Vimentin and N-cadherin levels were decreased in the cells, consistent with the Western blot results. Results were obtained from three independent experiments. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001. (**E**,**F**) Relative expression of vimentin and N-cadherin mRNA in the MDA-MB-231 cells were analyzed by real-time PCR. RNA levels are represented as fold increase relative to the level of the control (normalized to GAPDH mRNA level).

Real-time PCR (rt-PCR) showed that the mRNA level of E-cadherin was upregulated while the N-cadherin and Vimentin were downregulated significantly in response to actinomycin V treatment. These results confirmed that actinomycin V inhibited the epithelial-mesenchymal transition process in MDA-MB-231 cells.

### *2.5. Actinomycin V Suppresses the EMT Process by Reducing the Expression of SNAIL and SLUG*

EMT is a complex process which required a complicated network of transcription factors such as zinc finger and basic helix loop helix to participate in. Snail family transcriptional repressor 1 (also known as SNAIL) and Snail family transcriptional repressor 2 (SLUG) are proved to be the E-cadherin repressors and act as inducers of the invasion process when they are overexpressed in epithelial cell lines [18,19]. As shown in Figure 8, we found that actinomycin V treatment down-regulated the expression levels of snail and slug protein while the expression levels of zinc finger E-box binding homeobox 1 (ZEB1) and twist remained unchanged in MDA-MB-231 cells. Immunofluorescence staining also showed the same reduction effects after 1–2 nmol/L actinomycin V treatment. Therefore, we suggest the actinomycin V inhibitory effect on EMT process may through down-regulate the expression levels of the snail and slug protein.

**Figure 8.** Effects of actinomycin V on the expression levels of snail and slug. (**A**,**B**,**D**) MDA-MB-231 cells were treated with 0 to 2 nmol/L actinomycin V for 24 h then the protein expression of snail and slug was measured by Western blot. Results were obtained from three independent experiments. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001. (**C**) cells were treated with 0 to 2 nmol/L actinomycin V for 6 h and analyzed by snail and slug fluorescent signals. Cells were stained with anti-snail (red), slug (green) and DAPI (blue). Magnification ×200. Snail and slug levels were decreased in the cells, consistent with the Western blot results.

### **3. Discussion**

The majority reason of the high incidence and death rates of breast cancer among women is metastasis, which is initiated by the processes of adhesion, invasion and migration [20]. First, adhesion, it plays a vital role in maintaining cellular structure and the disruption of cellular adhesion may led to cell invasion and migration [6,7]. Cellular adhesion is mediated by a variety of membrane molecules, for example, cadherins, the Ca2+-dependent transmembrane proteins, is one of the major members of adhesion related membrane molecules [21]. More specially, reports have been confirmed that E-cadherin was required to initiate cell-cell adhesion and the procession of epithelial-mesenchymal transition (EMT) is closely related to its deletion [22,23]. In this paper, we found that the actinomycin V was effective in suppressing migration and invasion of breast cancer cells as revealed by morphological changes, wound-healing and Transwell assays. Moreover, actinomycin V treatment upregulated the protein and mRNA levels of E-cadherin, indicating that actinomycin V enhanced cell-cell adhesion in MDA-MB-231 cells. Next, we measured the N-cadherin and Vimentin, the other two important maker molecules of EMT, by Western blotting and immunofluorescent staining. E-cadherin serves as an invasion suppressor, N-cadherin, which is frequently upregulated in cancer cells, functions as an invasion promoter. The expression of N-cadherin induces epithelial cells transform the morphological into a fibroblastic phenotype and the biological function also changes to make epithelial cells more motile and invasive in the EMT process [24]. Vimentin is one of the major members of intermediate filament family and frequently overexpresses in a variety of cancer cells such as prostate cancer, breast cancer and lung cancer. The overexpression of vimentin in cancer cells promotes cellular proliferation, invasion and migration [25]. Our results showed that actinomycin V treatment significantly down-regulated both of the protein and mRNA expression levels of N-cadherin and vimentin, which may inhibit EMT in cells by reducing cells invasion and migration. These findings suggest that actinomycin V inhibits MDA-MB-231 cells migration and invasion via suppressing the EMT process.

A complicated network of transcription factors (TFs) are involved in EMT process. Factors from Snail, ZEB and helix-loop-helix families are acted as the inducers of epithelial-mesenchymal transition (EMT) and potent repressors of E-cadherin expression [26,27]. The activation of these factors initiates the EMT process. Then we examined the effects of actinomycin V on these family members including ZEB1, twist, slug, and snail. We found that the actinomycin V treatment down-regulated the expression levels of snail and slug protein while the expression levels of ZEB1 and twist remained unchanged. Immunofluorescence staining also showed us that the expression of snail and slug were reduced in MDA-MB-231 cells after actinomycin V treatment. It has been reported that snail and slug active during EMT and down-regulate the expression of adhesion genes like E-cadherin. Snail and slug also involved in the expression of the invasion promoter such as vimentin, N-cadherin, and fibronectin [27–30]. Therefore, the reduction of snail and slug may lead to the up-regulation of E-cadherin expression while the down-regulation of vimentin and N-cadherin and suppress the EMT process, which is consistent with the results showed in our studies. As a result, we conclude that the actinomycin V suppresses EMT process via down-regulating the snail family members snail and slug and subsequently inhibits the proliferation, migration, and invasion of breast cancer cells. On the other hand, it is noticed that the selectivity index of actinomycin V between cancer (such as MDA-MB-231) and noncancer cell lines (HMLE/MCF-10A) still remains low (around 4–5) as shown in Table 1. Toxicity investigation should be especially warranted in vitro/vivo for actinomycin V drug development.

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

### *4.1. Chemicals and Reagents*

Actinomycin V (>98%), actinomycin Xob (>98%) and actinomycin D (>98%) were provided by Dr. Xie (Shandong University, Weihai, China) isolated from *Streptomyces* sp. N1510.2 and identified by ESI-MS, 1H and 13C NMR data [31]. Compounds were dissolved in dimethylsulfoxide (DMSO) as 10 μmol/L stock solution and diluted according to experimental requirement. MTT (3-(4,5-Dimethylthiazol)-2,5-diphenyltetrazolium bromide) was purchased from Sigma-Aldrich Corp. (St. Louis, MO, USA). Antibodies against E-cadherin, N-cadherin, Vimentin, snail and ZEB1 were purchased from Cell Signaling Technology (CST, Inc., Beverly, MA, USA). Slug, twist, GAPDH antibodies and goat/donkey anti- mouse/rabbit IgG-Alexa Fluor 488/647 secondary antibodies were purchased from Abcam, Inc. (Cambridge, MA, USA). The primers used in this study were bought from Sangon Biotech Co Ltd. (Shanghai, China). All chemicals used in this study were commercial reagent grade products.

### *4.2. Cell Lines and Cell Culture*

Human breast cancer cell lines MCF-7, MDA-MB-231, BT474 and normal human breast epithelial cell lines HMLE and MCF-10A were purchased from the Shanghai Institute for Biological Sciences (SIBS), Chinese Academy of Sciences (Shanghai, China). MCF-7, MDA-MB-231 and BT474 were cultured in RPMI 1640 Medium (Hyclone Laboratories, Inc., Logan, UT, USA) and HMLE cells were cultured in DMEM/F-12 1:1 (Hyclone Laboratories, Inc.) containing 10% fetal bovine serum supplemented with 100 units/mL of penicillin and 100 μg/mL of streptomycin. MCF-10A cells were cultured in DMEM/F-12 1:1 containing insulin, hydrocortisone, EGF and 10% horse serum supplemented with 100 units/mL of penicillin and 100 μg/mL of streptomycin. All cell lines were cultured in a humidified incubator at 37 ◦C with 5% CO2.

### *4.3. MTT Assay*

Cell proliferation inhibitory effect of actinomycin V was evaluated by MTT assay. Cells were seed in 96-well plates at a density of 5 <sup>×</sup> 103 cells per well. After 24 h incubating, cells were treated with actinomycin V, actinomycin Xob and actinomycin D at various concentrations while adriamycin was used as the comparison. After 24–72 h of continuous culture, 15 μL MTT (5 g/L) was added to each well and incubated for another 4 h at 37 ◦C. The medium with MTT was removed and 150 μL per well of DMSO was added to dissolve the formazan. The absorbance of each well was measured by a microplate reader at 570 nm. IC50 values (concentration resulting in 50% inhibition of cell growth) for each cell line were calculated by plotting the untreated cells, which were considered to be 100% cell survival. Each text was performed in triplicate independently.

### *4.4. Wound-Healing Assay*

Cells in logarithmic growth phase were seeded into 6-well plates at a density of 3 <sup>×</sup> <sup>10</sup><sup>4</sup> cells per well. When the cells had adhered to 90%, they were scratched by micropipette tips vertically in the middle of each well. Each plate was washed with 3 mL PBS three times to remove the suspension cells. The cells in plate were starved for 12 h to eliminate the interference of proliferation in advance and then treated with 0 to 2 nmol/L actinomycin V for 24 h. The scratch width changes of each group were observed under a microscope, and the wound healing rate (%) was calculated as (1 − (scratch width of the actinomycin V group/scratch width of the control group)) × 100% to evaluate the migration ability of cells. Each experiment was performed in triplicate independently.

### *4.5. Transwell Assay*

We used transwell assays to measure the invasive capacity of cells. We took 50 μL Matrigel diluted with 1:8 of serum-free culture medium and added onto the upper chamber membrane of the insert. After solidifying into gel, cells were suspended in 200 μL serum-free culture medium at a destiny of <sup>1</sup> <sup>×</sup> 105 cells/mL and then added into the upper chamber of the insert, while the lower chamber were filled with 600 μL culture medium containing 20% FBS. Cells were treated with varying concentration of actinomycin V ranging from 0 to 2 nmol/L and incubated with 5% CO2 at 37 ◦C for 24 h. The medium in chamber was removed and the cells in the upper chamber were wiped by cotton swabs. Cells in lower chamber were fixed in formaldehyde for 20 min, then stained with 0.5% crystal violet for 15 min. After three times washing, five fields were taken randomly under a Nikon TE2000 microscope

(Nikon Instruments Inc., Tokyo, Japan) and the number of transmembrane cells were counted. Each experiment was performed in triplicate independently.

### *4.6. Immunofluorescence Staining*

Cells were seeded onto 12-mm round, glass over slips in 24-well plates and treated with 0 to 2 nmol/L actinomycin V for 6 h. Cells were washed and fixed with 4% Paraformaldehyde for 15 min, washed three times with cold PBS, and permeabilized in 0.1% Triton X-100 for 20 min at room temperature. To prevent non-specific antibody binding, the cells were washed and incubated with 3% goat serum for 30 min. Then cells were incubated with the E-cadherin, N-cadherin, vimentin, snail and slug antibody at 4 ◦C overnight, washed and incubated with donkey anti-rabbit IgG-Alexa Fluor 647 antibody or goat anti-mouse IgG-Alexa Fluor 488 antibody for 1 h. After washing three times, cells were counterstained with 4 μg/mL 4 ,6-diamidino-2-phenylindole (DAPI) for 10 min at room temperature. After washing three times with PBS-TX, cover slips containing the cells were then mounted using mounting medium (PBS:glycerol = 1:1 (*v*/*v*)). Fluorescence images were captured by using a fluorescence microscope.

### *4.7. Western Blot Analysis*

Cells were treated with indicated concentrations of actinomycin V for 24 h, then the protein from whole cell lysates was analyzed by Western blot assay. Samples were denatured by boiling with 1 × Laemmli buffer, subjected to 10% SDS-polyacrylamide gel electrophoresis and transferred to nitrocellulose membranes. The membranes were washed with distilled water and then blocked with 5% non-fat milk in TBS-T buffer (10 mmol/L Tris-HCl, 150 mmol/L NaCl and 0.05% Tween-20 (*v*/*v*), pH 7.8) for at least 1 h at room temperature. After a short wash in TBS-T buffer, the membranes were incubated in a solution containing monoclonal antibodies specific for GAPDH, Vimentin, E-cadherin, N-cadherin, snail, slug, twist and ZEB1 for 2 h at room temperature or overnight at 4 ◦C. Then, the membranes were washed with TBS-T buffer for 6 min three times and once in TBS buffer. Membranes were incubated in secondary HRP-conjugated goat anti-mouse IgG or donkey anti-rabbit IgG for 1 h at room temperature. Membranes were washed with TBS-T buffer three times and once in TBS buffer. Proteins on the membranes were visualized using the enhanced chemiluminescence detection system (ECL®, Amersham Biosciences). Proteins expression density values were quantified by gray analysis using the Image J 2.0 software (National Institutes of Health, Bethesda, MD, USA).

### *4.8. RNA Extraction and Relative Quantification by Real Time PCR*

Cells were seeded in 6-well plates and treated with 0 to 2 nmol/L actinomycin V for 24 h. Total RNA was extracted using RNeasy kit under the manufacturer's instructions and the purity of RNA was detected by OD260/280 of RNA sample (>1.8). Then we converted the RNA into cDNA using the ReverTra Aoe® qPCR RT Kit (QIAGEN, Hilden, Germany). Primers were designed by primer premier 5 for the E-cadherin gene (forward primer: 5 -CGAGAGCTACACGTTCACGG-3 and reverse primer: 5 -GGGTGTCGAGGGAAAAATAGG-3 ), N-cadherin gene (forward primer: 5 -CCTTTCAAACACAGCCACGG-3 and reverse primer: 5'-TGTTTGGGTCGGTCTGGATG-3'), and vimentin gene (forward primer: 5 -GACGCCATCAACACCGAGTT-3 and reverse primer: 5 -CTTTGTCGTTGGTTAGCTGGT-3 ). GAPDH gene (forward primer: 5 -CATCAAGAAGGTGGTGAAGCAGG-3 and reverse primer: 5 -TCAAAGGTGGAGGAGTGGGTGTCGC-3 ), was used as a control for the amount of RNA. Real-time PCR assay was used to detect the expression levels of the Vimentin, N-cadherin and E-cadherin genes and the amplification was performed in triplicate. For each reaction, we prepared 1×SYBR Green Realtime PCR Master Mix (TOYOBO, Osaka, Japan), 1 μL forward primer and reverse primer and 1 μg of template cDNA in a final volume of 20 μL. The condition of the amplification were 45 cycles of sequential denaturation (95 ◦C, 2 min), annealing (60 ◦C, 15 s), and extension (72 ◦C,

20 s). The ΔΔCT value was used to analyze the rt-qPCR data. All samples were measured in triplicate experiments independently.

### *4.9. Statistical Analysis*

Data are presented as Mean ± Standard deviation from triplicate experiments. All data were analyzed using One-way Analysis of Variance (ANOVA) for multiple comparisons by SPSS 16.0 (SPSS Inc., Chicago, IL, USA). Significant differences are indicated as follows: \* *p* < 0.05; \*\* *p* < 0.01; \*\*\* *p* < 0.001.

**Author Contributions:** Conceptualization, X.L.; Formal analysis, S.L.; Funding acquisition, X.L.; Investigation, S.L., C.Z., F.L., J.M., F.J., and Z.H.; Resources, W.X. and X.L.; Writing—original draft, S.L.; Writing—review & editing, S.L. and X.L.

**Funding:** This work was supported in part by grants from Shandong Provincial Natural Science Foundation (No. ZR2019MH001), the Program for Changjiang Scholars and Innovative Research Team in University (PCSIRT, No IRT\_17R68), the Fundamental Research Funds for the Central Universities (No.2019ZRJC004), and National Natural Science Foundation of China (No. 81273532).

**Acknowledgments:** The authors thank Weidong Xie (Shandong university, weihai) for providing actinomycin V, actinomycin Xob and actinomycin D.

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

### **References**

© 2019 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 (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Divergolides T–W with Apoptosis-Inducing Activity from the Mangrove-Derived Actinomycete** *Streptomyces* **sp. KFD18**

**Li-Man Zhou 1,2,†, Fan-Dong Kong 2,†, Qing-Yi Xie 2, Qing-Yun Ma 2, Zhong Hu 3, You-Xing Zhao 2,\* and Du-Qiang Luo 1,\***

Received: 19 March 2019; Accepted: 8 April 2019; Published: 11 April 2019

**Abstract:** Four new ansamycins, named divergolides T–W (**1**–**4**), along with two known analogs were isolated from the fermentation broth of the mangrove-derived actinomycete *Streptomyces* sp. KFD18. The structures of the compounds, including the absolute configurations of their stereogenic carbons, were determined by spectroscopic data and single-crystal X-ray diffraction analysis. Compounds **1**–**4** showed cytotoxic activity against the human gastric cancer cell line SGC-7901, the human leukemic cell line K562, the HeLa cell line, and the human lung carcinoma cell line A549, with **1** being the most active while compounds **5** and **6** were inactive against all the tested cell lines. Compounds **1** and **3** showed very potent and specific cytotoxic activities (IC50 2.8 and 4.7 μM, respectively) against the SGC-7901 cells. Further, the apoptosis-inducing effect of **1** and **3** against SGC-7901 cells was demonstrated by two kinds of staining methods for the first time.

**Keywords:** mangrove-derived actinomycete; ansamycins; divergolides; apoptosis-inducing activity

### **1. Introduction**