cribrochalina vasculum compounds activate intrinsic ......zovko et al., 2014 1 marine sponge...

Zovko et al., 2014

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Marine sponge Cribrochalina vasculum compounds activate intrinsic apoptotic signaling and inhibit

growth factor signaling cascades in non-small cell lung carcinoma

Ana Zovko1, Kristina Viktorsson

1*, Petra Hååg

1, Dimitry Kovalerchick

2, Katarina Färnegårdh

3,

Andrea Alimonti4,5

, Micha Ilan6, Shmuel Carmeli

2 and Rolf Lewensohn

1

1 Department of Oncology and Pathology, Karolinska Biomics Center, Karolinska Institutet,

Stockholm, Sweden, 2 School of Chemistry, Raymond and Beverly Sackler Faculty of Exact

Sciences, Tel Aviv University, Tel Aviv, Israel, 3 Science for Life Laboratory, Drug Discovery &

Development Platform, Department of Organic Chemistry, Stockholm University, Stockholm,

Sweden, 4 Molecular Oncology lab, Institute of Oncology Research (IOR), Bellinzona, Switzerland,

5 Atrahasis srl, Rome, Italy,

6 Department of Zoology, George S Wise Faculty of Life Sciences, Tel

Aviv University, Tel Aviv, Israel

Running title: C. vasculum compounds possess anti-tumor potential.

*Corresponding author: Kristina Viktorsson, Karolinska Biomics Center, Z5:01, Department of

Oncology-Pathology, Karolinska Institutet, SE-171 76 Stockholm, Sweden. Email:

[email protected], Phone: +46 8 517 701 77

on July 3, 2020. © 2014 American Association for Cancer Research. mct.aacrjournals.org Downloaded from

Author manuscripts have been peer reviewed and accepted for publication but have not yet been edited. Author Manuscript Published OnlineFirst on October 15, 2014; DOI: 10.1158/1535-7163.MCT-14-0329

mailto:[email protected]

http://mct.aacrjournals.org/

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Keywords: Sponge, small molecule, apoptosis, cytotoxicity, growth factor signaling, lung cancer

Conflicts of interest: The authors disclose no potential conflicts of interest.

Financial support

This study was supported by grants from the Swedish Cancer Society (R. Lewensohn grant

agreement 130550), Stockholm Cancer Society (K. Viktorsson grant agreement 131253, P. Hååg

131102 and R. Lewensohn 11 1213), Swedish Research Foundation (R. Lewensohn grant

agreement 90266701/2009), the Swedish National Board of Health and Welfare grant to R.

Lewensohn, the Stockholm County Council grant to R. Lewensohn, the Karolinska Institutet

Research Fund and the European Union (FP7/2007-2013) under grant agreement no KBBE-2010-

266033. A. Alimonti was supported by European Research Council starting grant (ERCsg) (GA

N°261342).




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Abstract

Marine derived compounds have been explored and considered as possible antitumor agents. In this

study we analyzed extracts of the sponge Cribrochalina vasculum for their ability to inhibit tumor

cell proliferation. Screening identified two acetylenic compounds of similar structure that showed

strong tumor specific toxicity in non small cell lung carcinoma (NSCLC) cells and small cell lung

carcinoma cells, less prominent toxicity in ovarian carcinoma while having no effect on normal

cells. These acetylenic compounds were found to cause a time dependent increase in activation of

apoptotic signaling involving cleavage of caspase-9, caspase-3 and PARP, as well as apoptotic cell

morphology in NSCLC cells, but not in normal fibroblasts. Further analysis demonstrated that these

compounds caused conformational change in Bak and Bax, resulted in loss of mitochondrial

potential and cytochrome c release in NSCLC cells. Moreover, a decreased phosphorylation of the

growth factor signaling kinases Akt, mTOR and ERK was evident and an increased phosphorylation

of JNK was observed. Thus these acetylenic compounds hold potential as novel therapeutic agents

that should be further explored for NSCLC and other tumor malignancies.




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Introduction

Marine organisms have been shown to be a great source of pharmaceutical compounds which can

be used for anticancer, antiviral or anti-inflammatory purposes. Since the 1970´s there is a steady

increase in the number of new compounds discovered and the number of registered patents is also

growing (1-3). Among the different marine organisms, sponges (Porifera) have been identified as

the most promising source of new compounds with anticancer potential (2-5). The leading role of

Porifera as a source of new anti-cancer compounds is likely attributed to their long evolutionary

history, extreme plasticity and their remarkably rich associated microbiota (4). Indeed many

sponge-derived compounds are at different stages of development (preclinical to phase III and pre-

registration) and also to some extent in clinical use (6, 7). Successful examples are discodermolide,

trabectedin, cytosar-U and eribulin (8-11). Cytosar-U more commonly known as Ara-C, identified

in Cryptotethia crypta, is clinically approved for the treatment of acute myeloid leukemia (12).

Another example is Eribulin, a truncated analogue of the natural product halichondrin-B isolated

from the sponge Halichondria okadai. This agent was approved by the U.S. Food and Drug

Administration in 2010, to treat patients with metastatic breast cancer (11). Trabectedin is yet

another example of a compound from the marine source, the Caribbean marine tunicate

Ecteinascidia turbinata, which has passed all steps of development into a pharmaceutical agent and

is approved for cancer patients with advanced or metastatic soft-tissue sarcoma or ovarian

carcinoma (10).

For non-small cell lung carcinoma (NSCLC), small cell lung carcinoma (SCLC) as well as ovarian

carcinoma prognosis is generally poor which in part is explained by the fact that these tumor cells

are highly resistant to known chemotherapeutic agents used in the disseminated stage of these




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diseases. For all these tumor entities failure to elicit apoptotic response as well as increased growth

factor signaling both contributes to the chemotherapy refractory phenotype (13, 14). Hence novel

agents, which can circumvent these signaling aberrations, are needed.

In the course of our screening for anti-tumor agent from marine sponges, (3S)-icos-4E-en-1-yn-3-ol

(1) and (3S)-14-methyldocos-4E-en-1-yn-3-ol (2) were isolated from Caribbean sponge

Cribrochalina vasculum (family Niphatidae, order Haplosclerida). Structure elucidation and

literature search of these compounds revealed that they both were previously described (15-17) and

reported to have anti-tumor activity, yet their mechanism of action for induction of tumor cell death

has not been described. Beside these compounds, several other related acetylenic alcohols have

been previously reported from the sponge C. vasculum. Such related acetylenic alcohols were

shown to exhibit immunosuppressive activity, toxicity toward brine shrimp and also to possess

antitumor activity (8, 15, 18, 19). Here we show that compound 1 causes strong cytotoxic activity in

tumor cells from NSCLC, SCLC and ovarian carcinoma but not in normal cell lines and primary

cells tested. Yet we see differences also among the tumor cells with NSCLC being more sensitive

than SCLC and ovarian carcinoma. For compound 2 a cytotoxic effect was evident in NSCLC and

SCLC whereas in ovarian carcinoma the difference compared to normal cells was less prominent.

Nevertheless, in NSCLC we can demonstrate that both these compounds, isolated from sponge C.

vasculum, activate intrinsic apoptotic signaling cascades i.e. activation of Bak/Bax, decreased

phosphorylation of Bad, depolarization of mitochondria, cytochrome c release from mitochondria

and activation of caspase 9 and caspase 3. Moreover, a decreased phosphorylation of Akt, mTOR

and ERK growth factor signaling kinases was evident and an increased phosphorylation of JNK was

observed. Thus compound 1, and to some extent compound 2, hold potential as therapeutic agents




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that should be further explored for different tumor malignancies with specific emphasis on NSCLC

and SCLC.

Materials and Methods

Collection and purification of extracts

Samples of the sponge C. vasculum were collected in Key Largo, Florida in June 2011. Further

details about the collection of the material can be found in Supplementary Materials and Methods,

section Collection of sponges. The freeze-dried sample (206 g), was extracted with 45:45:10

mixture of ethyl acetate:methanol:water to yield crude extract (43 g). Fractionations of crude extract

were guided by MTT cell viability assay (see Cell viability assay below) and each fraction was

tested for cell cytotoxicity in NSCLC U-1810 cells or in normal diploid fibroblasts WI-38 at 72 h of

continuous exposure. The fraction that gave the best therapeutic window was chosen for further

purification. Thus the crude extract was separated sequentially by different chromatographic

methods starting with reversed phase vacuum flash chromatography using a gradient of solvents,

from 100 % water through 100 % methanol and then 100 % ethyl acetate (Supplementary Fig.

S1A). The fraction 10 (90:10 mixture of methanol:water) obtained from the initial separation was

further separated twice on a size exclusion Sephadex LH-20 column using chloroform:methanol

(1:1 v/v) as mobile phase. In the last step of purification, the semi-pure fractions from the second

Sephadex LH-20 column (fractions 8-11) were combined and separated on a reversed phase semi-

preparative YMC C-8 HPLC column with a mixture 40:51:9 of acetonitrile:methanol:water as the

isocratic mobile phase. Several products were obtained from this last purification and pure

compounds (Supplementary Fig. S1B) were tested for tumor specific toxicity. Two pure substances,




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(3S)-icos-4E-en-1-yn-3-ol (1) (tR 41.8 min, 70.1 mg, 95% purity) and (3S)-14-methyldocos-4E-en-

1-yn-3-ol (2) (tR 67.0 min, 109.0 mg, 99% purity) were found to have the best therapeutic window

and were therefore chosen for further analysis.

The following characteristics were noted for compounds 1 and 2:

(3S)-icos-4E-en-1-yn-3-ol (1): colorless oil, [α]D 1.5 (c 0.35, CHCl3); UV (MeOH) () 204 (1400)

nm; IR (ATR) 3292, 2917, 2850 cm-1; For NMR data see Supplementary Table S1; GCMS SMB EI

m/z 292.3 M+, C20H38O (19).

(3S)-14-methyldocos-4E-en-1-yn-3-ol (2): Colorless oil; [α]D 6.4 (c 5.8, CHCl3); UV (MeOH) ()

204 (1300) nm; IR (ATR) 3303, 2922, 2852, 2098 cm-1; For NMR data see Supplementary Table

S2; GCMS SMB EI m/z 334.3 M+, C23H42O (19).

Cell culture and treatments

C. vasculum parental extract (fraction 10) and compounds 1 and 2 were diluted in DMSO to make

10 mg/ml stock solutions which were kept at -20 °C and diluted in cell culture media prior to use.

Data on stability of compound 2 after long term storage in DMSO and in cell culture media after 72

h incubation time at concentrations used in experiments can be found in Supplementary Materials

and Methods, sections Stability of compound 2 after long term storage and Stability of compound 2

in cell culture media.

The human SCLC U-1285 (20) and NSCLC U-1810 (21) were kind gifts from Uppsala University

where they were established and characterized (22). SCLC H69 and H82, ovarian cancer cell lines




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A2780 and SKOV-3, foreskin fibroblasts immortalized with hTERT BJ-5ta, bronchial epithelial

cells BEAS-2B and hTERT immortalized retina epithelial cells RPE-1 were all purchased from

ATCC (Mannassas, VA, USA). The lung fibroblast cell line WI-38 (23) was obtained from Coriell

Cell Line Repository (CCR, Camden, NJ, USA). Human cardiomyocyte primary cell culture was

acquired from Celprogen (Celprogen Inc., CA, USA). BJ-5ta was obtained in 2014, BEAS-2B and

cardiomyocytes in 2012, RPE-1 and SKOV-3 in 2011, A2780 in 2008 and U-1810, U-1285, H69

and H82 in 1996. Cell lines were authenticated by cell banks using the short tandem repeat

profiling. No authentication for these cell lines was done by the authors. Upon receipt cultures were

passaged for not more than 2 months and aliquots were frozen. For experiments each cell line was

thawed according to protocol and grown out for no more than 6 weeks for tumor and 3 for normal

cells before use in an experiment. Human peripheral blood mononuclear cells (PBMC) were

isolated from buffy coats of healthy donors (provided by the Department of Clinical Immunology

and Transfusion Medicine, Karolinska Hospital, Stockholm, Sweden) by using a standard Ficoll-

Hypaque gradient.

The U-1810, H69, H82, U-1285, A2780 cells and PBMC were cultured at 37 °C and 5 % CO2 in

RPMI-1640 medium (Sigma Aldrich, Stockholm, Sweden) containing 2 mM L-glutamine

(Invitrogen, Stockholm, Sweden) and 10 % heat-inactivated fetal bovine serum (FBS) (HyClone,

Täby, Sweden). The ovarian cancer cell line SKOV-3 was grown in McCoy´s 5A medium (Sigma

Aldrich) containing 2 mM L-glutamine and 10 % FBS. Lung fibroblasts WI-38 were maintained in

MEM (Sigma Aldrich) supplemented with 15 % FBS and 2 mM L-glutamine. RPE-1 (hTERT

immortalized retina epithelial cells) were cultured in DMEM/ F12 (Lonza, Belgium) with 10 % of

FBS. Cardiomyocytes were grown in human cardiomyocytes cell culture complete growth media




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with serum and antibiotics using Celprogen’s human cardiomyocyte cell culture extra-cellular

matrix pre-coated flasks (Celprogen Inc.). BJ-5ta were cultured in Dulbecco’s modified Eagle’s

medium (HyClone) supplemented with 2 mM L-glutamine and 10 % FBS. BEAS-2B were grown in

BEGM kit media (Lonza) and flasks were pre-coated with a mixture of 0.01mg/ml fibronectin

(Sigma Aldrich), 0.03 mg/ml bovine collagen type I glutamine (Invitrogen) and 0.01 mg/mL bovine

serum albumin (Sigma Aldrich) dissolved in BEBM (Lonza) to allow proper growth

Cell viability assay

The cytotoxicity induced by extracts or pure compounds was determined with the previously

described 3-(4,5-dimethylthiazol-2-yl)-2,5 diphenyltetrazolium bromide (MTT) assay (24) with

minor modifications. Cell viability experiments on tumor cells were carried out 24 h post seeding, at

a confluence of about 80 % whereas normal cells were completely confluent to mimic a non-

dividing state which is their natural state in the body. For all cells the amount of cells required were

titrated out in preparatory experiment and the following amount of cells were seeded in each well of

a 96 well plate in 90 μL of complete growth media: 5,000 cells (U-1810, H82, SKOV-3 cell lines

and cardiomyocytes), 7,500 cells (H69, A2780 and BJ-5ta), 10,000 cells (U-1285), 18,000 cells

(WI-38), 20,000 cells (BEAS-2B and RPE-1) and 400,000 (PBMC). All technical replicates were

done in triplicates and for each experiment the three biological replicates were carried out. Different

concentrations of parental extract and 1, 2 or equal volumes of DMSO (vol/vol) (negative control)

diluted in fresh medium were added into cells followed by post incubation for 24, 48 or 72 h. At the

end of the experiment, MTT solution (Sigma-Aldrich) was added (0.5 mg/ml) and the plates were

incubated for 4 h at 37 °C and 5 % CO2. Thereafter formed formazan crystals were dissolved in stop

solution (10 % SDS and 0.01 M HCl) and absorbance was measured at 595 nm. The resulting




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absorbance, which is proportional to the number of viable cells in each well, was recorded and %

cell survival was determined by comparing the absorbance in treated cells relative to the absorbance

in untreated cells which was set to 100 %. Values shown are the mean of three replicates in three

independent experiments ± SEM.

Analysis of apoptotic morphology and cytochrome c release

U-1810 and WI-38 cells were treated with 3 μM of 1, 2, 20 μM cisplatin or with equal volume of

DMSO and harvested after 24, 48 or 72 h. Approximately 40,000 cells was resuspended in 100 µl

PBS, cytospinned onto glass sides at 500 RPM for 4 min and fixed in 4 % paraformaldehyde (PFA)

for 5 min followed by ice cold acetone for 10 sec. Glass slides were dried and kept at -20 °C until

staining. To detect and quantify apoptotic morphology NSCLC U-1810 and WI-38 were stained

with 4,6′diamidino-2-phenylindole dihydrochloride (DAPI) (Vector Laboratories, CA, USA).

Apoptotic morphology was examined in a fluorescence microscope Zeiss Axioplan 2 imaging

microscope using Zeiss 403 lens and processed using the Axiovision software (Carl Zeiss

MicroImaging, Gottingen, Germany). The presence of fragmented nuclei was defined as apoptotic

feature and the percentage of such cells in total of 400 cells examined are presented. Cytochrome c

release was examined in U-1810 cells treated with either DMSO or 3 μM of 1 or 2 for 16 h.

Blocking was carried out with buffer (3 % BSA, 0.2 % TritonX100, 10 mM Hepes, pH 7.4) during

60 min at room temperature and the slides were subsequently incubated with primary antibody

against cytochrome c (1:200) (Cell signaling, Danvers, MA, USA) for 16 h at 4 °C, followed by 3

steps of washing in PBS and incubation with goat-anti rabbit alexa 488 secondary antibody

(Invitrogen). To visualize nuclei, slides were counterstained with DAPI (Vector Laboratories).

Staining was examined using a Zeiss Axioplan 2 imaging microscope with Zeiss 403 lens and




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processed using the Axiovision software (Carl Zeiss MicroImaging).

Mitochondrial permeability

The loss of mitochondrial membrane potential upon treatment with, DMSO, 3 µM of 1 or 2 for 24 h

was assessed by staining the cells with tetramethylrhodamine ethyl ester perchlorate (MitoPT™

TMRE; Immunochemistry Technologies Bloomington, MN, USA) as previously been described

(25). Briefly, cells were treated with, DMSO, 3 µM of 1 or 2 for 24 h. As a positive control for

depolarization of mitochondria the proton gradient uncoupling agent carbonyl cyanide 3-

chlorophenylhydrazone (CCCP) (50 μM, 45 min) was used. After harvesting, cells were stained

with TMRE (150 nM) for 20 min in the dark. Cells were centrifuged and resuspended in 1X Assay

buffer provided by the MitoPT™ TMRE staining kit. Cells (10,000/sample) were analyzed on the

FL-2 channel of Calibur flow cytometer (BD Bioscience, Franklin Lakes, NJ, USA). The

percentage of cells that lost their TMRE staining was gated and quantified and values shown are the

mean of three independent experiments ± SD.

Analysis of proapoptotic conformational changes of Bak and Bax

During apoptosis the pro-apoptotic Bcl-2 family proteins Bax and Bak undergo N-terminal

conformational changes leading to a pro-apoptotic state required for formation of pores in the

mitochondria and subsequent cytochrome c release (26, 27). To analyze N-terminal changes in Bak

or Bax in response to treatment NSCLC U-1810 cells were treated with DMSO or 3 µM DMSO

solution of 1 or 2 for 16 h. Cells were harvested and fixed in 4 % paraformaldehyde for 10 min and

washed in PBS. Fixed cells were stained with antibodies recognizing N-terminal activation-related

conformational changes in either Bak (AM03, clone TC100; Oncogene Research Products, CA,




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USA) or Bax (clone 6A7; BD Biosciences Pharmingen, San Diego, CA, USA). For staining cells

were resuspended in 100 μl of primary antibody diluted in PBS (Bak 1:50, Bax 1:250) containing

digitonin (100 μg/ml) as permeabilizing agent. After 1 h incubation at 4 ºC samples were washed in

PBS and incubated with secondary antibody Alexa 488 labeled anti-mouse antibody for 30 min at 4

ºC in the dark to enable monitoring by flow cytometry. Prior to FACS analysis samples were

washed in PBS and resuspended in 200 μl of PBS. The Bak or Bax associated IFL signal was

measured in the FL-1 channel of the FACS Calibur flow cytometer (BD Bioscience). Data was

processed using Cell Quest software (BD Bioscience) and change in fluorescence intensity was

determined by comparing percent of gated cells after treatment with that of untreated cells which

was set to 100 %. Data shown are the mean values of three independent experiments ± SD.

Western blot analysis

NSCLC U-1810 cells and WI-38 fibroblasts were seeded in 10 cm cell culture dishes at density of

1.2 million cells for U-1810 and 2 million for WI-38. After 24 h cells were treated for 4, 16, 24, 48

and 72 h with 3 µM 1 or 2 or with equal volume of DMSO. In experiment where apoptotic

induction was analyzed by PARP-1 and caspase cleavage, cells were treated for 24 h with

concentration of compounds that induce IC50 or IC70 in U-1810 cells or in WI-38.

Cells were harvested with Cell dissociation solution (Sigma Aldrich). Whole-cell lysates were

prepared in RIPA buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 0.5 % Igepal, 5 mM EDTA and

0.1 % SDS) supplemented with protease and phosphatase inhibitor cocktail tablets (Roche

Diagnostics AB, Stockholm, Sweden). Protein concentration was determined by BCA Protein

Assay Reagent (Interchim, Montiucon Cedex, France). 30-40 μg total protein of each extract were




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loaded to Bis Tris 4-12 % or Tris Acetate 3-8 % gels (NuPAGE, Invitrogen, Stockholm, Sweden)

and electrophoresed at 200V for 60 min in MES or MOPS running buffer (NuPAGE, Invitrogen).

Transfer to polyvinylidene fluoride (PVDF) membranes (Hybond-C Extra, Amersham Biosciences,

Piscataway, NJ, USA) was performed at 30V for 90 min in transfer buffer (NuPAGE, Invitrogen)

supplemented with 10 % methanol. After blocking nonspecific binding with the Odyssey blocking

buffer (Li-Cor Biosciences, Bad Homburg Germany) diluted in PBS (1:1) membranes were probed

with primary antibodies overnight at 4 °C and with secondary antibody for 1 h at room temperature.

Following primary antibodies diluted in blocking buffer were used: phospho-Akt (Thr308),

phospho-Akt (Ser473), Akt, phospho-Bad (Ser112), Bad, Bak, Bax, Bcl-xL, Caspase 3, Caspase 9,

phospho-mTOR (Ser2448), phospho-p44/42MAPK(Erk1/2) (Thr202/Tyr204),

p44/42MAPK(Erk1/2) (Cell Signaling, Danvers, MA, USA); JNK (FL), PARP (H250) (Santa Cruz

Biotechnology, Santa Cruz, CA, USA) and phospho-JNK1/JNK2 (Thr183/ Tyr185) (Abcam,

Cambridge, UK). Where indicated antibodies against β tubulin, β actin (Abcam, Cambridge, UK) or

GAPDH (Trevigen, Gaithersburg, MD, USA) were used as control of equal loading. IR-Dye-linked

secondary antibodies (LI-COR Biosciences, Bad Homburg, Germany) were used to image bands on

the Odyssey platform.

Cell cycle distribution

To assess cell cycle distribution, NSCLC U-1810 and WI-38 cells were treated with 1.3 M

(inducing IC50 in U-1810 at 24 h), 3.4, 6.8 and 16.2 μM (inducing IC50 in WI-38 at 24 h) of

compound 1 or an equal volume of DMSO for 24 h and fixed in 70 % ethanol. Cells were stained

with propidium iodide (PI) as described previously (28). Signals were recorded using FACS Calibur




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(BD Bioscience) and analyzed with ModFit LT (Verity Software House, Topsham, ME, USA).

Results

Isolation of anti-tumor extracts from Cribrochalina vasculum

Several compounds with antitumor activity have been isolated from marine derived organisms and

some of them are already in clinical use (8-11). Here we set out to isolate and characterize

antitumor compounds from the sponge C. vasculum with aim to reveal their antitumor mechanism

of action. For that purpose material was collected and extracted as described in Material and

Methods. Resulting fractions were tested for antitumor activity against NSCLC U-1810 cells and

for normal cell cytotoxicity using diploid fibroblasts WI-38 during 72 h continuous exposure.

Fraction 10 (named parental extract) from the C. vasculum extract induced significant cytotoxicity

of NSCLC cells and almost completely inhibited cell growth at a concentration of 0.5 μg/ml while

only minor cytotoxicity was evident in a normal diploid fibroblasts WI-38 up to a concentration of

25 μg/ml (Fig. 1A-B). Given the observed NSCLC cell specific cytotoxicity we continued the

purification of this fraction as outlined in Material and Methods and Supplementary Fig. S1A.

Chemical structure elucidation of Cribrochalina vasculum anti-tumor active fractions

identifies (3S)-icos-4E-en-1-yn-3-ol (1) and (3S)-14-methyldocos-4E-en-1-yn-3-ol (2)

HPLC fractionation of the parental extract from C. vasculum (Supplementary Fig. S1B) yielded two

known alkyl-4-en-1-yn-3-ol derivatives; 70.1 mg (0.035 % of sponge dry weight) of 95 % pure

(3S)-icos-4E-en-1-yn-3-ol (1) and 109 mg (0.055 % of sponge dry weight) of 99 % pure (3S)-14-

methyldocos-4E-en-1-yn-3-ol (2) (19). The structures of 1 and 2 were confirmed by 1D and 2D




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NMR techniques. EIMS and MS/MS techniques were used to elucidate the position of the branched

methyl substituent in compound 2 (Supplementary Fig. S1C) (19). The absolute stereochemistry of

1 and 2 was determined by the Mosher method (29) since their optical rotations were small.

Structural analysis showed that compounds 1 and 2 are structurally related acetylenic alcohols both

containing a C4-C5 trans double bond and the same absolute configuration of the hydroxyl group.

The structure determination also showed that the compounds differed in the length of their carbon

chains and that 2 has a branched methyl substituent at position 14 (Fig. 1C).

Cribrochalina vasculum compounds 1 and 2 have tumor specific cytotoxicity

We next continued with the acetylene containing compounds 1 and 2 to reveal their antitumor effect

and delineate their mechanism of action. Similar to the parental extract compounds 1 and 2 induced

toxicity in NSCLC U-1810 cells but not in normal diploid fibroblasts WI-38 until 3 μM was used

(Fig. 2A). A clear cytotoxicity was also evident when treating the NSCLC U-1810 cells with

different concentrations of 1 and 2 for 72 h and examining cell viability with MTT assay (Fig. 2B).

Thus, at concentration of 3 μM 1 reduced U-1810 cell viability for about 90 %, and 2 for 60 %.

Importantly, applying the same concentration of these compounds to WI-38 fibroblasts caused little

or no cytoxicity suggesting a tumor selective window for these compounds. Thus the concentration

of compound 1 and 2 needed to inhibit 50 % of cell viability (IC50) was more than 10 times higher

for WI-38 than for the U-1810 cell line (Supplementary Table S4). Next, the effect of 1 and 2 on

cell cytotoxicity was examined also at 24 h and 48 h post treatment in U-1810 and WI-38

(Supplementary Fig. S2). A clear time dependent increase in the cytotoxic effect of both compounds

toward U-1810 cells was evident with a higher effect seen on the later time point. In contrast,

neither 1 nor 2 impaired survival of diploid fibroblasts WI-38 at these earlier time points at doses up




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to 15 μM and 30 μM, respectively (Supplementary Fig. S2 and Supplementary Table S3).

Cribrochalina vasculum compounds 1 and 2 induce prominent cytotoxicity in SCLC but not in

normal cells

Next we extended our analyses onto three small cell lung carcinoma cell lines (U-1285, H69 and

H82) and two ovarian cell lines (SKOV-3 and A2780) (Fig. 2C, Supplementary Table S4). In order

to asses normal cell toxicity human cardiomyocytes, human peripheral blood mononuclear cells

(PBMC), foreskin fibroblasts immortalized with hTERT (BJ-5ta), bronchial epithelial cells (BEAS-

2B) and hTERT immortalized retina epithelial cells (RPE-1) were used in the analyses (Fig. 2D,

Supplementary Table S4).

Of the SCLC cell lines, H82 showed the highest sensitivity towards compound 1 with an IC50 of 1.1

µM and with an almost complete inhibition of cell survival at 1.5 µM (Fig. 2C, left panel). The

toxicity of compound 1 was slightly lower for U-1285 and H69, yet at 3 μM both these cell lines

showed an 80 % decrease in cell viability and the corresponding IC50 values were 1.6 and 2.2 μM,

respectively (Supplementary Table S4). For compound 2 U-1285, H69 and H82 responded with

about 80 % decrease in cell viability upon treatment with 3 µM concentration (Fig. 2C, right panel).

IC50 values were 1.8 µM for U-1285, 1.3 µM for H69 and 1.1 µM for H82 (Supplementary Table

S4). In ovarian carcinoma cells compound 1 caused a similar 80 % reduction in cell viability at 3

μM as seen in SCLC whereas for compound 2 both A2780 and SKOV-3 cells were less responsive

(Fig. 2C). Compound 1 reached IC50 in A2780 at 1.8 μM and in SKOV-3 at 2.1 μM. Toxicity of

compound 2 toward ovarian carcinoma was not so prominent, IC50 was reached only at 3.4 µM in

A2780 and 4.9 µM in SKOV-3 cell line (Supplementary Table S4). Taken together compound 1 and




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2 both demonstrate toxicity toward SCLC while for ovarian carcinoma compound 1 have a greater

potency than compound 2.

Both compounds were tested against several normal cells. Treatment with 3 μM concentration of

compound 1, which caused high cytotoxicity in all tumor cell lines tested, induced only 20 %

reduction in cell viability in cardiomyocytes and BEAS-2B whereas in PBCM and BJ-5ta about 40

% cytotoxicity was evident (Fig. 2 D, left panel). With the applied concentrations IC50 value

couldn’t be reached for cardiomyocytes and BJ-5ta whereas for RPE-1 it was 6.6 μM, PBMC 7.8

μM and for BEAS-2B 12.9 μM (Supplementary Table S4). Compound 2 at 3 μM, a concentration

that caused at least a 50% cell cytotoxicity in LC induced less than 15% cytotoxicity in

cardiomyocytes, BJ-5ta, BEAS-2B and RPE-1 whereas in PBMC about a 35 % cytotoxic effect was

evident (Fig. 2 D, right panel). With compound 2 IC50 value was reached only for PBCM and RPE-

1 (8.5 μM for PBMC and 10.4 for RPE-1) (Supplementary Table S4).

In summary, the cytotoxic effect of compound 1 was significantly lower in all normal cells than in

tumor cells whereas for compound 2 the tumor-selective effect was less prominent but evident in

NSCLC and SCLC.

Cribrochalina vasculum compounds 1 and 2 induce apoptosis in NSCLC tumor cells but not in

diploid fibroblasts

In order to further understand mechanism of action of these C. vasculum compounds, their effect on

apoptosis induction was examined in NSCLC cells (Supplementary Fig. S3A and Fig. 3A).

Treatment of NSCLC U-1810 cells with either 1 or 2 (3 µM, 48 h) caused a prominent induction of

apoptotic morphology of the cell nuclei whereas no major alteration was observed in diploid




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fibroblasts WI-38 upon treatment (Supplementary Fig. S3A). Induction of apoptotic morphology in

U-1810 cells was also quantified at 24-72 h post treatment with 1 or 2 (3 μM) (Fig. 3A). Already

after 24 h 1 induced apoptotic morphology in about 30 % of the cells with a further increase after 48

h and with 80 % of the cells showing apoptotic nuclear morphology at 72 h post treatment (Fig.

3A). Compound 2 induced apoptosis was delayed as compared to 1 but nevertheless 48 h post

treatment about 40 % of the U-1810 cells had fragmented nuclei and at 72 h almost 60 % showed a

prominent apoptotic response (Fig. 3A). Both compounds induced less than 2 % apoptosis in

fibroblasts treated for 72 h (data not shown). Cisplatin which was used as a positive control induced

apoptotic morphology in 26 % of U-1810 cells showing that these cells are capable to undergo

apoptosis.

Cribrochalina vasculum compound 1 induces G2 arrest in NSCLC but not in normal diploid

fibroblasts

Loss of proliferation capacity and induction of cytotoxicity may also be attributed to inhibition of

cell cycle progression. The effect of 1 on cell cycle distribution was therefore also evaluated (Fig.

3B). NSCLC U-1810 or WI-38 fibroblasts were treated with different concentrations of 1 for 24 h.

In NSCLC U-1810 cells, treatment with 1 was found to induce a significant dose dependent

increase in the number of cells arrested in the G2/M phase (Fig. 3B, left graph). In contrast, cell

cycle distribution of normal fibroblasts showed no alteration after treatment with compound 1 even

at IC50 concentration (Fig. 3B, right graph). Thus the results showed that compound 1 clearly

induces cell cycle arrest in G2/M in NSCLC tumor cells and this is a tumor-specific effect.

Cribrochalina vasculum compounds 1 and 2 induce mitochondrial depolarization, activate




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19

caspases-9 and 3 and cause PARP cleavage in NSCLC cells but not normal diploid fibroblasts

One important gateway to apoptosis is mitochondrial depolarization allowing cytochrome c to be

released to the cytosol where it subsequently causes the pro-caspase-9 to be activated within the

apoptosome complex. Caspase-9 in turn triggers cleavage of pro-caspase-3/7 into active proteases

thereby giving rise to apoptotic morphology (30). The effect of 1 and 2 on depolarization of

mitochondria and release of cytochrome c was therefore examined. NSCLC U-1810 cells were

treated with compounds 1 or 2 (3 μM) for 24 h and mitochondrial depolarization was examined

with TMRE (Fig. 4A). Upon membrane depolarization, the potential of mitochondria is lost and the

TMRE dye leaks into cytosol. Thus cells which have a decrease in TMRE staining represent cells

with depolarized mitochondria and these cells were quantified. The mitochondrial uncoupler CCCP

(50 μM, 45 min) was used as positive control showing that TMRE is capable of detecting

alterations in mitochondrial potential (Fig. 4A). Importantly, treatment with either compounds

resulted in prominent mitochondrial depolarization (Fig. 4A). Hence both compounds were capable

of disrupting mitochondrial membrane potential.

Next the release of cytochrome c after treatment of NSCLC U-1810 cells with 1 and 2 (3 μM) for

16 h was analyzed by immunofluorescence staining (Fig. 4B). A punctate staining pattern of

cytochrome c indicative of mitochondrial localization was found in untreated cells whereas upon

treatment with 1 or 2 diffuse staining pattern indicating release of cytochrome from mitochondria to

cytosol was evident (Fig. 4B).

In order to study whether depolarization of mitochondria and release of cytochrome c in response to

1 or 2 also resulted in the downstream activation of caspase 9/3 and subsequent apoptotic-associated




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20

cleavage of the caspase-3 substrate PARP-1 NSCLC U-1810 cells were treated with 1 or 2 for 24 h

at a concentration that caused 50 % and 70 % of cell death respectively (Fig. 4C). As can be seen in

U-1810 cells both caspase-9 and caspase-3 were cleaved into active forms (35-37 and 15-17 kDa

respectively) upon treatment with IC50 or IC70 of either compound. Treatment also resulted in clear

cleavage of PARP-1 into the pro-apoptotic associated 89 kDa fragment confirming that these

compounds indeed triggered an apoptotic response (Fig. 4C). Normal fibroblasts WI-38 were used

for comparison and treated with same concentrations as U-1810 cells. Importantly under these

conditions no caspase-3 or PARP-1 cleavage was evident in normal WI-38 fibroblasts (Fig. 4C).

However the lower pro-apoptotic activity of compounds observed in fibroblasts could be due to cell

permeability rather than to a difference in actual mode of cell kill. In order to see if these

compounds at all were able to induce apoptosis in these normal lung fibroblasts, concentrations that

induced 50 % and 70 % cell kill at 24 h was applied. Even at these concentrations, no PARP-1 or

caspases cleavage was observed in the fibroblast further demonstrating a NSCLC cell specific pro-

apoptotic activity of these compounds (Fig. 4D).

We also examined the kinetic of caspases and PARP-1 cleavage after treatment with 3 μM of 2 for

24, 48 and 72 h (Supplementary Fig. S3B). In line with MTT and nuclear apoptotic morphology

data, cleaved versus full-length caspase-3, caspase-9 and PARP-1 all clearly showed an increase

over time in response to 2. Thus there is a clear NSCLC specific activation of apoptosis by both

compounds.

Cribrochalina vasculum compounds 1 and 2 trigger activation of Bak and Bax and JNK

signaling and inhibit PI3K/Akt and MAPK ERK survival signaling in NSCLC cells




Zovko et al., 2014

21

Activation of the Bcl-2 proteins Bak and Bax is one important gateway for the intrinsic

mitochondrial pathway of apoptosis (26, 27). The effect of compound 1 and 2 on Bax and Bak

conformational changes associated with activation was therefore analyzed (Fig. 5A and 5B).

Indeed, treatment of NSCLC U-1810 cells with 3 μM of 1 or 2 resulted in prominent activation of

Bak indicated by a shift of histogram peak to the right (Fig. 5A and 5B right). An activation of Bax

was also observed after treatment of NSCLC U-1810 cells with either 3 μM of 1 or 2, yet less

pronounced than for Bak (Fig. 5A and 5B left). The total expression of Bax and Bak was also

examined using conformation-independent antibodies in western blot (Fig. 5C). However, no

changes in their expression levels were evident upon treatment indicating that the observed

Bak/Bax activation upon 1 or 2 relates to N-terminal conformational changes of these proteins.

The antiapoptotic Bcl-xL protein is reported to inhibit Bak/Bax complex formation and subsequent

cytochrome c release (31). The expression level of Bcl-xL was therefore examined after treatment

with compound 1 or 2 and a clear decrease in Bcl-xL expression was evident after treatment with 1,

while no change was observed after treatment with 2 (Fig. 5C). All in all, these results show that

both 1 and 2 can activate the Bak/Bax rheostat in NSCLC cells which is in line with the observed

depolarization of mitochondria, release of cytochrome c and activation of caspases via the intrinsic

route after treatment with these compounds.

Bak and Bax complex formation is in part regulated by the BH3-only protein Bad which upon

phosphorylation is sequestered by binding to 14-3-3 protein preventing its pro-apoptotic action (32).

The effect of 1 or 2 in causing dephosphorylation of Bad was therefore examined. Indeed a

decreased phosphorylation of Bad at Ser112 was clearly evident in response to treatment with 1

already at 16 h post treatment and still evident at 24 h in NSCLC U-1810 cells (Fig. 5C). In




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22

response to treatment with 2 a moderate decrease in Bad phosphorylation was observed only at 24 h

(Fig. 5C). The observed alteration on Bad Ser112 in response to 1 was confirmed to be a result of

altered signaling as no difference in total Bad protein expression was evident upon treatment (Fig.

5C). Thus both compounds partly inhibit Bad phosphorylation indicating a putative pro-apoptotic

action of Bad at mitochondria mediated apoptotic signaling in NSCLC cells.

Several growth factor signaling cascades are reported to control Bad phosphorylation thereby

inhibiting its pro-apoptotic signaling capacity. The effect of 1 and 2 on the phosphorylation status of

the Bad-regulating kinases Akt, Erk1/2 as well as on JNK, a kinase shown to promote apoptotic

signaling (13, 33) was therefore evaluated. A decreased phosphorylation of Akt at both Ser473 and

Thr308 was evident in NSCLC U-1810 cells after treatment with compound 1 or compound 2 with

an earlier response seen with 1 (Fig. 5D). Deactivation of mTOR was also evident after 1 as the

phosphorylation on Ser2448 declined (Fig. 5D). These data suggest that 1 and 2 impair PI3K/AKT

pathway signaling and may in this way inhibit proliferation and survival of NSCLC cells. A

decreased phosphorylation on Thr202/Tyr304 of ERK1/2 was also evident after treatment with both

1 and 2 again with a more prominent deactivation caused by compound 1 (Fig. 5D). Hence a

decreased pro-survival signaling via the MAPK ERK may also contribute to the observed cytotoxic

activity of these two compounds in NSCLC cells. A sustained increase in JNK phosphorylation is

reported to be critical for efficient induction of apoptotic signaling (33). In line with this 1 caused

increased phosphorylation of JNK1/JNK2 at 16 h and 24 h whereas in response to 2 a similar

increase in phosphorylation was seen only at 24 h in these NSCLC cells. Thus this increased JNK

phosphorylation may in part contribute to the observed apoptotic signaling in response to these

compounds in this tumor type.




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23

Discussion

Here we analyzed the ability of C. vasculum derived metabolites to inhibit proliferation, induce

cytotoxicity and trigger apoptotic signaling in NSCLC, SCLC and ovarian carcinoma. It has

previously been shown that compounds isolated from this sponge can kill tumor cells of different

origin albeit through unknown mechanism of action (19). We show here that the two acetylenic

compounds 1 and 2 as well as the parental mixture of these compounds from which they were

separated, indeed cause tumor-specific cytotoxicity in NSCLC and SCLC. Importantly, we

demonstrate that for both compounds this cytotoxicity is tumor specific, as all normal cells tested

representing heart tissue, bronchial and retina epithelium, normal peripheral blood mononuclear

cells, and foreskin or lung fibroblasts remained unaffected at concentrations that were toxic for

NSCLC and SCLC. In ovarian carcinoma cells compound 1 but not compound 2 caused tumor-

specific cell death. For compound 2 a cytotoxic effect was evident in NSCLC and SCLC whereas in

ovarian carcinoma the difference compared to normal cells was less prominent.

With respect to NSCLC cytotoxicity we for the first time show that compounds from C. vasculum

can induce a prominent apoptotic response whereas in normal fibroblasts no induction of apoptosis

was seen after exposure to the same concentration of 1 or 2. Moreover no apoptosis was seen in

normal fibroblasts even when concentrations that caused 50 % or 70 % cytotoxic response in cell

viability assay were used. This indicates that the induction of apoptosis is a tumor selective path

that might be a consequence of inhibition of cell survival signaling pathways which tumor but not

normal cells rely upon on for protection against apoptotic cell death.

At an early stage of the intrinsic apoptotic pathway the pro-apoptotic Bcl-2 family proteins Bak and




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24

Bax are activated leading to permeabilization of the mitochondrial membrane, release of

cytochrome c and activation of caspases within the apoptosome (34, 35). We demonstrate that both

1 and 2 causes all these events in NSCLC cells. However, a more prominent activation of Bak over

Bax was evident in these NSCLC cells. This is likely attributed to the basic signaling propensity of

these NSCLC cells rather than specific action of these compounds to activation of Bak

preferentially over Bax. This conclusion is based on the fact that we showed similar less

pronounced activation of Bax also after treatment with other agents e.g. cisplatin and radiation in

these NSCLC cells (25).

When comparing 1 and 2 with respect to Bak activation and mitochondria-mediated apoptotic

signaling, compound 2 was found to be less efficient. Compound 1 is, at the applied concentration 3

µM, more toxic toward U-1810 cells than is equimolar concentration of compound 2, likely

explaining the more pronounced Bak and Bax activation and cytochrome c release. There was also

the less efficient decrease of Bcl-xL protein expression upon exposure to 3 µM of compound 2. The

presence of Bcl-xL could sequester Bak in an inactive state thereby preventing cytochrome c release

and proper intrinsic route to caspase activation (34). We demonstrate that both 1 and 2 subsequently

trigger the cleavage and activation of caspase-9 and thereafter also caspase-3 showing that a full

apoptotic signaling cascade is active in these NSCLC tumor cells but not in normal diploid

fibroblasts. At their IC50 and IC70 values both compounds are equally efficient in induction of PARP

and caspases cleavage suggesting that they have similar potency in apoptosis induction. Altogether,

our results demonstrate that both C. vasculum 1 and 2 induce cell death by activation of intrinsic

apoptotic pathway and this effect is tumor cell specific. One of the regulators of Bax and Bak

activation is the BH3 only protein Bad which antagonize the antiapoptotic proteins Bcl-2 and Bcl-xl




Zovko et al., 2014

25

enabling Bak/Bax complex formation and cytochrome c release (36). Bad function as an integrator

of multiple growth factor signaling cascades including the PI3K/Akt and the Raf/MEK/ERK

pathway which antagonize Bad function by phosphorylation causing its binding and sequestration to

14-3-3 protein (37, 38). Akt inactivates Bad by phosphorylation at site Ser136 (39) while MAPK

signaling cascade inactivates Bad protein at Ser112 and Ser155 (40-43). With respect to 1 and 2

Bad phosphorylation at Ser112 was markedly inhibited indicating a role for MAPK signaling

pathway while the basal level of phosphorylation of the other sites i.e. in untreated cells were weak

making effect of treatment unsecure (data not shown). Given this altered phosphorylation of Bad

Ser112 by compound 1 and 2 we next examined how multiple growth and apoptosis promoting

kinases in NSCLC cells were affected. A decrease in ERK phosphorylation was observed which is

in line with the decrease in Bad protein phosphorylated at Ser112. Although results for inactivation

of Bad protein at site Ser136 were inconclusive a major impairment of phosphorylation of Akt at

Ser473 and Thr308 upon treatment with these compounds was evident. Deactivation of Akt is

reported to cause activation of pro-apoptotic proteins, cause alterations in mitochondrial membrane

potential, release of cytochrome c and caspase activation leading to activation of apoptosis (37, 43,

44). Hence the impaired Akt phosphorylation upon treatment with these compounds fits well with

observed apoptotic changes in these NSCLC cells.

We and others have linked NSCLC apoptotic response to a sustained JNK activation (33, 45-49). In

line with this and the observed apoptotic response, phosphorylation of JNK was increased by these

C. vasculum derived compounds. All in all, our data clearly demonstrate that 1 and 2 inhibit

proliferation and induce apoptosis through regulation of Ras/Raf/MAP kinase pathway as well as

deactivation of Akt signaling in NSCLC cells. As both Akt and MAPK kinase signaling are




Zovko et al., 2014

26

controlled by growth factor receptors these results may indicate that a common growth factor

receptor instrumental in controlling their activity could indeed be a target of C. vasculum

compounds. Further studies are however needed to reveal if that is the case. We observed that Bak

and Bax activation, cytochrome c release, downregulation of Bcl-xL, Bad, Akt, mTor,

p43/p44MAPK (ERK1/2), JNK1/JNK2 phosphorylation alteration were delayed or less affected by

2 than by 1 in these NSCLC cells when used at equimolar concentrations. Yet given that 1 has a

higher toxicity than 2 one cannot rule out that this is the underlying effect. Therefore at this point

we can only say that both compounds are capable of inducing apoptosis in tumor cells.

We show that 1 is also capable of enhancing cytotoxicity through induction of G2/M cell cycle

arrest in tumor cells. It has been shown that Akt regulates cell cycle by regulation of cyclins and

CDK inhibitors (50-53). In addition JNK is involved in cell cycle progression and mediates G2/M

phase cell cycle arrest in different tumor cells (54-59). As both; decreased phosphorylation of Akt

and increased phosphorylation of JNK is evident after treatment with 1, these signaling cascades

may be responsible for the observed G2/M arrest upon treatment. Further studies are however

required to uncover if this is the case.

Albeit compounds 1 and 2 only show structural variance in the length of their carbon chains and the

methyl substituent at position 14, a great difference in their potency is evident when it comes to

induction of cytotoxicity in NSCLC, SCLC and more importantly in ovarian carcinoma where 1 but

not 2 shows tumor-specific cytotoxicity. The cytotoxic response by 1 was much more prominent in

NSCLC and also appeared at an earlier time point. One possible reason for the less prominent

effects observed in tumor cells, especially in ovarian carcinoma, by compound 2 could be lower

affinity of drug to target a growth factor receptor upstream of these pathways. Yet another




Zovko et al., 2014

27

possibility is that their target upstream of these kinases are different and with the compound 2

targeting a signaling component which play a less prominent role in ovarian carcinoma.

Permeability of different cell types for the different compounds could also clearly have a role.

Hence further research is required in order to put an answer to these issues.

In conclusion our data reveal that 1 isolated from marine sponge C. vasculum causes tumor specific

cell death in NSCLC, SCLC and ovarian carcinoma. For compound 2 we found such an activity to

be evident in NSCLC and SCLC but not in ovarian carcinoma. Our findings are summarized in Fig.

6. We show that in NSCLC these compounds trigger a prominent response of the intrinsic apoptotic

pathway (activation of Bak/Bax, decreased phosphorylation of Bad, depolarization of mitochondria,

release of cytochrome c and activation of caspase-9 and caspase-3) and cell cycle arrest in G2/M

phase with a concomitant inhibition of both Akt and ERK/MAPK growth promoting pathways.

Moreover, we show that in NSCLC these compounds trigger a sustained JNK activation which may

contribute to the observed pro-apoptotic capacity. Taken together our results therefore implicate

that one or several growth factor receptors could be target of 1 and 2 either directly or indirectly and

in this way trigger cell death through the intrinsic apoptotic pathway and cell cycle arrest. Further

studies of the effects of compounds on growth factor receptors are therefore warranted in order to

reveal mechanism of action, develop these compounds into suitable pharmacological leads as well

as to understand their role in treatment of NSCLC and SCLC but also other tumor malignancies.

Acknowledgments

We are grateful to Mrs Birgitta Mörk and Miss Therese Juntti for excellent technical assistance.




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Figure Legends

Figure 1. Cribrochalina vasculum semi-pure fraction shows specific tumor cell cytotoxicity

and generates two pure compounds. (A) Fraction 10 [crude mixture of (3S)-alkyl-4E-en-1-yn-3-

ol, named parental] after three fractionation steps of sponge Cribrochalina vasculum extract was

tested for capacity to induce tumor specific cytotoxicity in NSCLC U-1810 cells or in diploid

fibroblasts WI-38 after 72 h of continuous exposure to indicated concentrations (phase contrast

images, 10x magnification, the scale bar equals 0.5 mm). (B) Cells were treated as in (A) and cell

viability was examined using MTT cell viability assay. Cell viability is given as % of cell survival

as compared to DMSO solvent treated cells. Data shown is the mean of three independent

experiments ±SEM. (C) Structure of (3S)-icos-4E-en-1-yn-3-ol (1) and (3S)-14-methyldocos-4E-

en-1-yn-3-ol (2).

Figure 2. Pure compounds isolated from Cribrochalina vasculum extract demonstrate high

tumor selective cytotoxicity in NSCLC and SCLC cells. (A) Either of the two isolated

compounds (1 and 2) at concentration 3 μM or equal volume of DMSO solvent was added to

NSCLC U-1810 and normal diploid fibroblasts WI-38 for 72 h. Phase contrast pictures showing

significant toxicity in tumor but not normal cells (magnification 20x, the scale bar equals 0.25 mm).

(B) The compounds 1 and 2 were profiled for their cytotoxic capacity in the same cells as in (A) at

72 h of continuous exposure using MTT cell viability assay. Cell viability is given as % of cell

survival as compared to DMSO solvent treated cells. Data shown is the mean of three independent

experiments ±SEM. The IC30, IC50 and IC70 concentrations are given in Supplementary Table S3.

(C) The cytotoxicity of 1 and 2 after 72 h of treatment was examined in additional tumor cell lines:

SCLC (U- 1285, H69 and H82) and ovarian cell lines (A2780 and SKOV-3). Normal diploid




Zovko et al., 2014

36

fibroblasts (WI-38) were included for comparison. Cell viability was examined as in (B). The IC30,

IC50 and IC70 concentrations are given in Supplementary Table S4. (D) The cytotoxicity of 1 and 2

after 72 h of treatment was examined in normal cells: human peripheral blood mononuclear cells

(PBMC), bronchial epithelial cells (BEAS-2B), human cardiomyocytes, hTERT immortalized retina

epithelial cells (RPE-1) and foreskin fibroblasts immortalized with hTERT (BJ-5ta). Normal diploid

fibroblasts (WI-38) were included for comparison. Cell viability is given as in (B). The IC30, IC50

and IC70 concentrations are given in Supplementary Table S4.

Figure 3. Compounds 1 and 2 induce apoptosis and cell cycle arrest in NSCLC cells but not in

diploid fibroblasts. (A) NSCLC U-1810 cells or diploid fibroblasts WI-38 were treated with

DMSO, 1 or 2 (3 μM) for 24, 48 and 72 h. Cisplatin (20 μM, 72 h) treated U-1810 cells were used

as a positive control for induction of apoptotic morphology. Nuclear morphology of cells was

examined after fixation in 4 % paraformaldehyde and staining with DAPI. Pictures showing nuclear

morphology are given in Supplementary Fig. S3A. The percentage of cells displaying apoptotic

morphology was quantified in 400 cells in three independent experiments using fluorescence

microscopy. Data shown is the mean and bars represent SD, *p<0.05. (B) Flow cytometric

examination of the cell cycle distribution in U-1810 (left) and WI-38 (right) cells treated with 1.3

M, 3.4, 6.8 and 16.2 μM concentration of 1 or equal volume of DMSO for 24 h. The percentage of

cells in G1 phase, S phase, and G2/M phase are given. Data shown is the mean of three independent

experiments ±SD, * P < 0.001

Figure 4. Compounds 1 and 2 induce depolarization of mitochondria, cytochrome c release

and activation of caspases in NSCLC cells but not in diploid fibroblasts. (A) NSCLC U-1810

cells were treated with 3 μM 1 or 2 for 24 h. CCCP (50 μM) was added to cells 45 min before




Zovko et al., 2014

37

staining as positive control. Depolarization of mitochondria was examined by staining with 150 nM

TMRE for 20 min in media. Left: The signal was recorded in the FL-2 channel on FACS calibur.

Histogram showing TMRE-associated fluorescence. Filled grey: DMSO; Solid line: 1 or 2 treated

cells. Right: Quantification of cells that lost their TMRE signals (i.e. depolarized mitochondria).

Data shown is the mean with bars representing SD, p<0.05. (B) Cytochrome c release was

examined by immunofluorescence staining of U-1810 cells after treatment with either DMSO or 3

μM of 1 or 2 for 16 h (magnification 100x, the scale bar equals 15μm). Blue: DAPI staining of

nucleus; Green: Cytochrome c. (C) NSCLC U-1810 cells or diploid fibroblasts WI-38 were treated

with compound 1 or 2 at concentrations that induced IC50 and IC70 in U-1810 cells at 24 h.

Cleavage of caspase-3, caspase-9 or PARP-1 was examined after 24 h. β tubulin was used to

visualize equal loading. (D) Diploid fibroblasts WI-38 were treated with 1 or 2 at concentrations

that at 24 h inhibited growth to 50% or 70 % (i.e. IC50 and IC70) respectively. Cleavage of caspase-

3, caspase-9 or PARP-1 was examined after 24 h. β tubulin was used to visualize equal loading.

Figure 5. Compounds 1 and 2 activate proapoptotic Bcl-2 family members and JNK signaling,

decrease Bad and Akt phosphorylation and inhibit MAPK ERK survival signaling in NSCLC

cells. (A) Conformational changes in Bax and Bak in response to 1 or 2 were examined in NSCLC

U-1810. Histograms showing Bax and Bak activation. Filled grey: DMSO; Solid line: 1 or 2 treated

cells. (B) Quantification of Bax (left) and Bak (right) associated IFL. Data shown is the mean of

three independent experiments ±SD. (C) NSCLC U-1810 cells were treated with 3 μM of either 1

or 2 for 4, 16 and 24 h or with equal volume of DMSO for 24 h and Bcl-xL, Bax, Bak and Bad

expression and Bad phosphorylation at Ser112 were examined by western blot. GAPDH or β

tubulin served as loading controls. (D) Deactivation of AKT on phosphoresidues Ser473, Thr308,

mTOR Ser2448, p42/44MAPK(Erk1/2) on Thr202/Tyr204 and JNK1/JNK2 on Thr183/Tyr185




Zovko et al., 2014

38

and total forms of AKT, p42/44MAPK(Erk1/2) ERK, JNK1/JNK2 were examined at 4, 16 and 24 h

post addition of 1 or 2 or 24 h treatment with equal volume of DMSO. GAPDH or β actin were used

as loading controls.

Figure 6. Schematic illustration of the proposed molecular mechanism of C. vasculum

compounds in NSCLC cells. Experimental data from NSCLC cells supports that compounds 1 and

2 indirectly or directly target growth factor receptors thereby interfering with downstream

proliferative signaling pathways Akt and MAPK i.e. decrease phosphorylation of ERK and Akt.

Both compounds were found to deactivate Akt signaling leading to impaired Bad phosphorylation

and activation of pro-apoptotic proteins Bak and Bax. Compounds were also found to trigger

depolarization of mitochondria, release of cytochrome c and activation of caspases. Compound 1

was also found to inhibit progression of cell cycle by arresting NSCLC cells in G2/M phase.




A B

0 1 10 25

µg/ml

NSCLC U-1810

Diploid fibroblasts WI-38

C

Parental [µg/ml]

% C

ell v

iability

DMSO 0.5 1 5 10 25 500

20

40

60

80

100

120

NSCLC U-1810

Diploid fibroblast WI-38

Figure 1




B

C

D

A

DMSO Compound 1

Diploid

fibroblasts

WI-38

NSCLC

U-1810

(3 µM)Compound 2

(3 µM)

Compound 1 [µM]

% c

ell

su

rviv

al

0 1 2 3 4 50

20

40

60

80

100

120

10 15 20

BEAS-2B CardiomyocytesPBMC

BJ-5ta Diploid fibroblasts WI-38RPE-1

Compound 2 [µM]

% c

ell

su

rviv

al

0 1 2 3 4 50

20

40

60

80

100

120

10 15 20

Compound [µM]

% c

ell s

urv

iva

l

0 1 2 30

20

40

60

80

100

120

4 6 8 10 12 1520

NSCLCU-1810

Diploid fibroblastsWI-38

Compound 2 Compound 2Compound 1 Compound 1

Figure 2

Compound 1 [µM]

% c

ell s

urv

iva

l

0 1 2 3 40

20

40

60

80

100

120

6 8 17

OC A2780SCLC U-1285 SCLC H69 SCLC H82 OC SKOV-3


Compound 2 [µM]

% c

ell

surv

ival

0 1 2 3 4 5 60

20

40

60

80

100

120

8 10 15




A

B NSCLC

U-1810

% C

ells

0

25

50

75

100

G1

S

G2/M

DMSO 1.3 3.4 6.8 16.2 µM


% C

ells

0

25

50

75

100

DMSO 1.3 3.4 6.8 16.2 µM

Hours post treatment

% A

po

pto

tic c

ells

24 48 720

20

40

60

80

100Compound 2

DMSOCompound 1

*

*

* *

*

*

*

*

* * *

* * *

Figure 3




AC

ou

nts

FL 2 TMRE

TMRE: CCCP

TMRE: Compound 1

TMRE: Compound 2DMSO Compound 1

(3 µΜ)B (3µM)

(3 µM)

Compound 2

C D

Diploid fibroblasts

WI-38

NSCLC

U-1810

DMSO IC₅₀ IC₇₀ IC₅₀ IC₇₀

Caspase-3 (FL)

PARP-1 (FL)

PARP-1 (CL)

35 kDa

17 kDa15 kDa

50 kDa

116 kDa

89 kDa

50 kDa

Caspase-3 (CL)

tubulinβ

β

Comp. 1 Comp. 2

DMSO IC₅₀ IC₇₀ IC₅₀ IC₇₀

Comp. 1 Comp. 2

Diploid fibroblasts

WI-38

IC₅₀ IC₇₀ IC₅₀ IC₇₀

Comp. 1 Comp. 2

tubulin

DMSO

35 kDa

17 kDa15 kDa

50 kDa

116 kDa

89 kDa

50 kDa

Caspase-3 (FL)

PARP-1 (FL)

PARP-1 (CL)

Caspase-3 (CL)

tubulinβ

β

tubulin

47 kDa

37 kDa

35 kDa

47 kDa

37 kDa

35 kDa

Caspase-9 (FL)

Caspase-9 (CL)

Caspase-9 (FL)

Caspase-9 (CL)

(In U-1810) (In WI-38)

Fo

ld T

MR

E s

tain

ing

0.0

0.5

1.0

**

*

DMSO CCCP Comp 1 Comp 2

(3 µM)

Figure 4




A

phospho-Bad (Ser112)

Bcl-xL

FL Bax/Bak

Co

unts

C

GAPDH

GAPDH

B

HOURS DMSO 4 16 24

Compound 1

4 16 24

Bax: Compound 1

(3µM)

Bax: Compound 2 (3µM)

Bak: Compound 1

(3µM)

Bak: Compound 2 (3µM)

Compound 2

phospho-Akt (Ser473)

phospho-Akt (Thr308)

Akt

phospho- p42/44MAPK(Erk1/2) (Thr202/Tyr204)

actin

p42/44MAPK(Erk1/2)

phospho-JNK1/JNK2 (Thr183/Tyr185)

GAPDH

phospho-mTOR (Ser2448)

JNK

β

β

actin

actinβ

GAPDH

D

HOURS DMSO 4 16 24

Compound 1

4 16 24

Compound 2

actinβ

actinβ

Bad

βtubulin

Bax

βtubulin

Bak

βtubulin

Fo

ld F

luo

rescence Inte

nsity B

ax

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

4.5

5.0

5.5

6.0p= 0.002

p=0.02

Comp. 1 Comp. 2DMSO

Fo

ld F

luo

rescence Inte

nsity B

ak

2468

10121416182022242628 p= 0.009

p=0.00002

Comp. 1 Comp. 2DMSO

BAX BAK

βtubulin

Figure 5




Figure 6




Published OnlineFirst October 15, 2014.Mol Cancer Ther Ana Zovko, Kristina Viktorsson, Petra Hååg, et al. signaling cascades in non-small cell lung carcinomaintrinsic apoptotic signaling and inhibit growth factor

compounds activateCribrochalina vasculumMarine sponge

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