Cell fusions in mammals

Cell fusions are important to fertilization, placentation, development of skeletal muscle and bone, calcium homeostasis and the immune defense system. Additionally, cell fusions participate in tissue repair and may be important to cancer development and progression. A large number of factors appear to regulate cell fusions, including receptors and ligands, membrane domain organizing proteins, proteases, signaling molecules and fusogenic proteins forming alpha-helical bundles that bring membranes close together. The syncytin family of proteins represent true fusogens and the founding member, syncytin-1, has been documented to be involved in fusions between placental trophoblasts, between cancer cells and between cancer cells and host cells. We review the literature with emphasis on the syncytin family and propose that syncytins may represent universal fusogens in primates and rodents, which work together with a number of other proteins to regulate the cell fusion machinery.
Cell fusions in the developing mammal
The fusion between the sperm and the egg marks the beginning of a new individual and ensuing development of either man or beast depends upon a number of additional cell fusion events (reviewed by Ogle et al. 2005). Thus, mononuclear cytotrophoblasts fuse to form the syncytiotrophoblast layer of the placenta in man and certain other mammals. Syncytiotrophoblasts control the exchange of gases, nutrients and waste products between the fetus and the mother, protect the fetus against the maternal immune system and are also responsible for the production of hormones, which like chorionic gonadotropin, regulate the continuation of pregnancy. In addition, in the developing individual, myoblasts fuse to form multinucleated skeletal muscle fibers, while cells of monocytic origin fuse to form osteoclasts, which participate in bone sculpturing and remodeling as well as in the regulation of serum calcium concentrations.
Cell fusions contribute to tissue repair
It is well-known that skeletal muscle is regenerated through fusion of muscle fibers with satellite cells (Bischoff 1994) and that macrophages may fuse to form multinucleated giant cells with enhanced phagocytic capabilities in response to injury and antigenic challenges (Vignery 2005). Recent data indicate that additional cell fusions may contribute to tissue repair in the adult (Vassilopoulos et al. 2003; Wang et al. 2003). Bone-marrow-derived (BMD) cells have been shown to fuse with hepatic cells, nerve cells and gastrointestinal cells and the theory has been put forward that such fusions may serve to repair damaged or corrupted cells (Alvarez-Dolado et al. 2003; Rizvi et al. 2006; Vassilopoulos et al.2003; Wang et al. 2003). However, it is still debated whether such fusions are important to tissue repair and whether they engage BMD stem cells or more differentiated fusogenic cells like monocytes/macrophages (reviewed by Vignery 2005). Intriguingly, the accumulation of macrophages in injured organs may reflect not only the need to remove debris but also to repair compromised cells through heterotypic fusions (Vignery 2005). Moreover, in recent transplant experiments, irradiated mice showed evidence of fusions between crypt stem cells and donor BMD cells (Rizvi et al. 2006). In thus transplanted mice, representatives of all cell lineages derived from intestinal crypt stem cells (absorptive columnar, enteroendocrine, goblet and Paneth cells) showed expression of chromosomal markers characterizing donor BMD cells. This phenomenon appeared to depend upon tissue injury since expression of donor cell markers never was detected in non-irradiated mice (Rizvi et al. 2006). An alternative to fusions has been suggested by Holmgren et al. (1999) and de la Taille et al. (1999), who have demonstrated that cells phagocytosing apoptotic cells may acquire functional DNA. Moreover, a mechanism of cell–cell invasion (entosis) was recently described and could potentially transfer DNA between cells (Overholtzer et al. 2007). Thus, viral as well as normal sequences from apoptotic cells have been found to be active also in phagocytosing cells (Holmgren et al. 1999). This observation challenges our understanding of apoptosis as an efficient way of clearing corrupted or alien DNA.
Cell fusions during viral infections
Infections with enveloped viruses such as HIV-1 may also lead to cell fusions. The viral genome encodes envelope (Env) proteins, which bind to cell surface receptors and assist the virus in entering the cell. The infected cell commences to synthesize Env proteins, which, upon insertion in the plasmalemma, engages receptor proteins in neighboring cells and initiate fusions. Interestingly, modifications of the cytoplasmic tail of several Env proteins appear to modify fusions. Thus, proteolytic cleavage of the cytoplasmic tail regulates fusions and also cellular signaling events, such as tyrosine kinase activity, may be involved in regulation of virally induced cell fusions (Kubo et al. 2003).
Receptor proteins may bind Env proteins from different classes of viruses. Thus, a cell infected by a specific virus becomes resistant to infections with other viruses that bind to the same receptor. This phenomenon, which probably reflects receptor saturation, has led to the grouping of retroviruses into different subtypes, which bind to the same receptor. Other types of viruses encode fusogenic proteins (like influenza hemagglutinin), which do not bind to surface receptors, but which liberate a fusion peptide in the acidic milieu of endosomes and thus initiate viral–host cell fusion following endocytic uptake of viral particles. Both these proteins and the Env proteins are referred to as class I viral fusion proteins and contain hydrophobic fusion peptides buried within their sequences. Following receptor engagement or endosomal acidification the fusion peptides are liberated by proteolytic cleavage. The cleaved peptides undergo conformational changes, resulting in hairpin-like, alpha-helical bundles, which bring the viral and cell membranes into close apposition and thereby facilitates fusion (reviewed by Chen and Olson 2005).
Cancer–host cell fusions
In 1911 Aichel suggested that cancer cells might spontaneously fuse with host cells to produce hybrids with supernumerary chromosomes that could evolve into cells of increased malignancy. Subsequent studies have confirmed this prediction by demonstrating that cells with mixed phenotypes spontaneously appear in co-cultures of normal and malignant cells (Barski and Cornefert 1962; Busund et al. 2002; Mortensen et al. 2004; Wakeling et al. 1994) (Figs. 1, 2, 3). Additionally, transplanted tumor cells of animal (Busund et al. 2003; Chakraborty et al. 2000; Fortuna et al. 1990; Kerbel et al. 1983; Larizza et al. 1984a; Wiener et al. 1974a, b) or human (Goldenberg et al. 1974; Goldenberg and Pavia 1982; Mortensen et al. 2004) origin may fuse with and acquire phenotypic characteristics of normal host cells (Fig. 4). In some of the in vivo studies, data suggested that the putative fusion partner was of macrophage or endothelial origin but, in most cases, definite identification was not achieved. In a few cases, it has also been demonstrated that the fused cells expressed genetic markers of both parental cell types (Goldenberg et al. 1974; Mortensen et al. 2004; Wiener et al. 1974a, b) (Figs. 2, 3). Mortensen et al. (2004) demonstrated that, initially after fusion, bi- or multinucleated cells formed and that such cells had the parental genomes seggregated in diffferent nuclei (heterokaryons) (Fig. 2a–e). Subsequently, mitotic figures appeared showing an admixture of the parental chromosomes (Fig. 2f–j). Eventually, cells with the parental genomes mixed in a single nucleus (synkaryons) were detected (Fig. 2k–o). In agreement with the notion that synkaryons appear after mitotic divison of heterokaryons, synkaryons are usually detected in pairs (Fig. 3). A few recent studies present compelling evidence that cells with genetic characteristics of hybrid cells also may appear in human tumors (Andersen et al. 2007; Chakraborty et al. 2004; Yilmaz et al. 2005; Streubel et al. 2004). Thus, in two cases of renal carcinomas, arising in bone marrow transplant recipients, some tumor cells showed genetic markers characterizing the healthy donor (Chakraborty et al. 2004; Yilmaz et al. 2005). Moreover, in patients with multiple myelomas, osteoclasts have been shown to contain an admixture of nuclei, of which some possess tumor-specific chromosomal translocations while others are devoid of translocations (Andersen et al. 2007). Finally, lymphoma-specific genetic abnormalities were described in endothelial cells in B cell lymphomas (Streubel et al. 2004). Although these data do not formally prove that tumor–host cell hybrids did form in these patients, the combined results from studies in vitro and in vivo do present compelling evidence that cell fusions do occur in tumors (reviewed by Pawelek 2005).
Fig. 1Cancer–endothelial cell fusion. Co-culture of human breast cancer (MCF-7) cells and human umbilical vein endothelial cells (HUVEC). Cells were stained for the cancer cell marker cytokeratin (red, c), the endothelial cell marker vimentin (green, b) and DNA (bisbenzimide, blue, d). Note in the merged image (a) one fused, multinucleated cell reacting for both cytokeratin and vimentin (orange-yellow)Fig. 2Chromosomal markers and cell fusions. Co-cultures of human breast cancer cells and bovine endothelial cells. The cultures were subjected to fluorescent in situ hybridization (FISH) with probes recognizing all bovine chromosomes (green; b, g, l) and all human chromosomes (red; c, h, m) and counter-stained for DNA with DAPI (blue; d, i, n) and observed in differential interference contrast (DIC; e, j, o). Merged red-green images are shown in a, f, k. Note that initially, binuclear cells (heterokaryons) form, having one bovine and one human nucleus (a–e). Occasionally, tri- or multinuclear cells with different admixtures of bovine and human nuclei are also detected. At longer times after mixing, mitotic figures, containing an admixture of bovine and human chromosomes appear (f–j) and, eventually, cells with a single nucleus, containing an admixture of the two genomes in mixture (synkaryons) are detected (k–o)Fig. 3Chromosomal markers and cell fusions. Mixed culture of human breast cancer cells and bovine endothelial cells submitted to double FISH as in Fig. 2 (a double FISH; b combined DAPI and DIC). Low power micrograph showing a pair of synkaryons with the bovine and human genomes admixed in single nuclei (arrows). In addition, nuclei hybridizing only for the bovine (green) or only for the human (red) genome occurFig. 4Cancer–host cell fusion in vivo. a, b Section from lung of a nude mice injected with human breast cancer (MDA-MB-231) cells in the tail vein (Mortensen et al. 2004). The section underwent FISH for the mouse genome (red) and the human genome (green) (a) and DNA was counterstained with DAPI (b; blue). Note one nucleus in which the human and mouse genome co-localize (arrow). C: Similar section, stained with an antibody detecting human (but not mouse) p53 (red; p53 is mutated and overexpressed by the breast cancer cells), an antibody to beta-catenin (to mark cell membranes) and for DNA with bisbenzimide (blue). Note a micrometastasis of human breast cancer cells having violet (red + blue) fluorescent nuclei. d Section stained for human p53 (red) and the endothelial marker von Willebrand factor (green) and DNA (blue). Note a human cancer cell with a violet (red + blue) nucleus showing membrane-staining for von Willebrand factor. Since von Willebrand factor is not normally expressed by the breast cancer cells, this image is suggestive of a fusion between a human breast cancer cell and a mouse endothelial cell. Similar results were obtained using double FISH for the human and mouse genome and imunofluorescent staining for von Willebrand factor (described by Mortensen et al. 2004)
The essential question regarding cancer–host cell fusions is, of course, if they are relevant to the patient. In fact, there are two opposing views. The first is based on early experiments on fusions induced to occur between cancer cells and normal cells in culture. With few exceptions, such experiments revealed that malignancy was suppressed (Harris et al. 1969; Harris 1988; Stanbridge 1976; Wiener et al. 1974a, b). These studies were, in fact, seminal to the discovery of tumor suppressor genes (reviewed by Anderson and Stanbridge 1993). Since tumor suppressor genes, like p53 and Rb, frequently are inactivated in cancer cells, fusions would present cancer cells with unperturbed tumor suppressors from the normal fusion partner and consequently initiate cell cycle arrest or apoptosis. Although, this certainly applied for the cell types studied in the contributions cited above, it may not be a general rule. Thus, production of monoclonal antibodies depends upon the fact that it is possible to fuse antibody-producing spleen cells with myeloma cells to obtain hybridomas that retain the unlimited proliferative ability of the tumor cell partner and the antibody production of the normal cell (Kohler and Milstein 1975). In fact, several studies documented that some fusions may lead to cells of increased malignancy (Barski and Cornefert 1962; Busund et al. 2003; Chakraborty et al. 2001; De Baetselier et al. 1981; Kerbel et al. 1983; Larizza et al. 1984b; Pawelek 2000; Rachkovsky et al. 1998). Possibly, the genetic make-up of tumors may dictate the outcome of individual cancer–host cell fusions. Moreover, fusions lead to aneuploidy and chromosomal instability, which characterizes most cancers and may, by itself, stimulate carcinogenesis (Duelli et al. 2007).
Mechanisms behind cell–cell fusions
Fusions between normal cells
Cell fusion events must be extremely well controlled. Due to their major importance to fertilization, placentation, muscle development, bone structure, calcium homeostasis and the immune defense system, much effort has gone into elucidating mechanisms underlying cell–cell fusions. Additionally, the potential role of cell fusions as a repair mechanism and the role of cell fusions in cancer development and progression have further stimulated research in this field. In spite of this, much less is known about cell–cell fusion mechanisms than is known about how intracellular membranes fuse through v- and t-SNAREs. Interestingly, engineered flipping of the v- and t-SNARE machinery has been shown to promote cell–cell fusions (Weber et al. 1998). However, it is evident that this mechanism is not a physiological mediator of cell–cell fusions. Interestingly, v- and t-SNAREs act similar to class I viral fusion proteins in that they form bundles of alpha-helices, which result in membrane apposition and fusion (Blumenthal et al. 2003; Jahn et al. 2003). Of the many proteins, which to date have been shown (or proposed) to be involved in cell fusions in mammals, only the syncytin family appears to use a similar alpha-helical mechanism. That they do is not surprising in view of the fact that syncytins represent conserved endogenous retroviral Env sequences. The founding family member, syncytin-1, was discovered as a protein capable of mediating fusions between cytotrophoblasts into syncytiotrophoblasts (Blond et al. 2000; Mi et al. 2000). This capability may have contributed to a high degree of evolutionary conservation of the syncytin-1 sequence. Syncytin-1 represents the Env protein of the human endogenous retroviral (HERV) W sequence, which entered the primate genome 25–40 million years ago. In contrast, most other retroviral sequences inserted in our genome have been subject to inactivating changes and probably represent garbage sequences. Molecular studies have shown that syncytin-1 (Env W) shares a structure similar to class I viral fusion proteins, especially in the region of the N- and C-terminal heptad repeats (NHR and CHR), and shares a common fusion mechanism with these proteins (Chang et al. 2004; Gong et al. 2005). A synthetic peptide derived from the CHR is also capable of inhibiting syncytin-1-mediated fusions by perturbing this mechanism (Chang et al. 2004). Syncytin-1 binds to the D-type retroviral receptor ASCT-2 (Blond et al. 2000) and may use another neutral amino acid transporter (ASCT-1) as an auxiliary receptor (Lavillette et al. 2000). Syncytin antibodies, syncytin-1 downregulation through antisense oligonucleotides and the syncytin-1 CHR peptide have been shown to inhibit fusions between trophoblast-derived cells (Blond et al. 2000; Chang et al. 2004; Mi et al. 2000). Agents increasing cellular levels of cAMP or cAMP analogues have been shown to promote cytotrophoblast fusions in vitro (Keryer et al. 1998) and such agents are also known to elevate protein and mRNA levels of syncytin-1 in isolated cytotrophoblasts (Frendo et al. 2003). Also estradiol may regulate syncytin-1 expression (Carino et al. 2003) and the placenta-specific transcription factor GCMa interacts with two upstream sites in the HERV-W 5′-long terminal repeat and stimulates syncytin-1 transcription (Yu et al. 2002). Additionally, experimentally induced truncations in the cytoplasmic tail of syncytin-1 increases its fusogenicity (Drewlo et al. 2006), similar to what has been observed for some virally derived Env proteins (reviewed by Kubo et al. 2003). However, if modifications in the cytoplasmic tail of syncytin-1 are of physiological importance for regulating fusogenicity has yet to be demonstrated.
A second syncytin-family member, syncytin-2, was subsequently found also to be expressed in the placenta (Blaise et al. 2003). Syncytin-2 also represents a highly conserved endogenous retroviral envelope gene and is derived from the HERV FRD sequence (Blaise et al. 2003). Immunocytochemical studies have localized syncytin-1 primarily to syncytiotrophoblasts as well as to cytotrophoblasts (Fig. 5). The exact localization is somewhat controversial (reviewed by Potgens et al. 2004), which may reflect differences in antibodies, fixation procedures and controls. A major point of concern is also the degree of cross-reactivity to the related sequences in syncytin-2, which recently was localized to a subpopulation of cytotrophoblasts (Malassiné et al. 2008). The syncytin-1 receptor protein ASCT-2 was recently localized to cytotrophoblasts (Hayward et al. 2007). More studies on the exact distribution of syncytin expression in the placenta using syncytin-1 and -2 specific antibodies seem warranted in order to exactly localize where fusions are likely to occur.
Fig. 5Human term placenta immunocytochemically stained with a syncytin-1 peptide antiserum. Staining occurs in the syncytiotrophoblasts, which predominate at this stage. Nuclei are lightly counterstained with haematoxylin
Trophoblast cell fusions are, however, not unique to the primate placenta. Thus, also in the mouse placenta, trophoblasts fuse to form syncytiotrophoblasts but mice do, of course, not express HERV sequences. Amazingly, an in-silico search of the mouse genome unraveled the existence of two murine endogenous retroviral (MERV) Env genes, labeled syncytin-A and -B (Dupresssoir et al. 2005). These genes were also fusogenic and orthologous genes were present in additional species of muridae (rats, gerbils, voles and hamsters) and appear to have entered the rodent lineage some 20 million years ago (Dupresssoir et al. 2005). They were expressed in the placenta and at least syncytin-A has been shown to be involved in the formation of syncytiotrophoblasts in mice (Gong et al. 2007). This represents an amazing example of parallel acquisition of retroviral genes of importance to reproduction in primates and rodents and poses the question whether also other species may have acquired similar viral genes of importance to cell fusions. The ability of syncytins to induce cell fusions may not be their only physiologic role. Thus, recent studies have shown that while syncytin-2 and syncytin-B also possess immunosuppressive activity, syncytin-1 and syncytin-A do not (Mangeney et al. 2007). Syncytin-1 expression was originally detected in the placenta and testis (Blond et al. 2000; Mi et al. 2000) but subsequent studies have revealed the presence of syncytin-1 also in the brain (Antony et al. 2004) and in breast, colon and endometrial cancers (Bjerregaard et al. 2006; Strick et al. 2007, Larsen, Talts, Andersen, Bjerregaard and Larsson: work in progress). Syncytin-1 may also regulate production of inflammatory mediators (Antony et al. 2004).
A number of additional molecules have been shown to be important to cell fusions. In fact, in all systems studied so far, from mating yeast to man, a bewildering array of mechanisms have been identified (see Chen and Olson 2005; Chen et al. 2007; Oren-Suissa and Podbilewicz 2007 for recent reviews). Molecules potentially involved in mammalian cell fusions include ADAM (a disintegrin and metalloproteinase domain) 12 (meltrin alpha), which has been associated with myoblast and osteoclast cell fusions (Abe et al. 1999; Galliano et al. 2000; Gilpin et al. 1998; Yagami-Hiromasa et al. 1995). In addition, ADAMs 1 and 2 (fertilins) may be involved in sperm–oocyte fusions, but do not seem indispensable for this function and, in man, the fertilin alpha gene is dysfunctional (Chen and Olson 2005; Cho et al. 1997, 1998, Evans et al. 1998; Jury et al. 1997, 1998). In contrast, the tetraspanin protein CD9 is needed for sperm–oocyte fusions (Le Naour et al. 2000). CD9 is also expressed by BeWo trophoblast tumor (choriocarcinoma) cells and has been linked both to BeWo invasiveness and to invasion during mouse embryo implantation (Hirano et al. 1999; Liu et al. 2006). CD9 has, together with another tetraspanin family member, CD81, also been linked to myoblast fusion and myotube maintenance (Tachibana and Hemler 1999). Moreover, antibodies to either CD9 or CD81 have been shown to block fusions induced by Mason—Pfizer monkey virus—a D-type retrovirus (Duelli et al. 2005). Tetraspanins are known to organize other proteins into membrane microdomains and may link to the actin cytoskeleton via EWI (Glu-Trp-Ile) and ERM (ezrin–radixin–moesin) proteins (Hemler 2003; Sala-Valdés et al. 2006). Possibly, their role as organizers of other proteins into microdomains may play a role in their involvement with cell fusions (Zivyat et al. 2006). They do not express characteristics of fusogenic proteins like the syncytins, SNAREs and class I viral envelope proteins, but are coexpressed with syncytin-1, at least in BeWo cells. A macrophage fusion receptor (MFR, SIRPalpha), resembling CD4–the cell surface receptor for HIV—has been identified in macrophages (Saginario et al. 1995, 1998; van den Berg et al. 2005). MFR, which belongs to the immunoglobulin superfamily, binds another member of this family, CD47. CD47 is also structurally related to proteins expressed by Vaccinia and Variola viruses (Chen et al. 2007). Whereas expression of MFR is restricted to myeloid cells and neurons, CD47 is ubiquitously expressed. MFR is transiently induced in macrophages at the onset of fusion while CD47 expression is constant. It has been hypothesized that CD47 initially binds to a long form of MFR to secure recognition and then switches to bind a shorter form to bring the plasma membranes closer (5–10 nm) for fusion (Vignery 2005). Additionally, or alternatively, CD47 may promote calcium entry by forming a membrane pore (reviewed in Chen et al. 2007). Also the hyaluronan receptor, CD44, is induced transiently when macrophages start to fuse. CD44 and MFR are subsequently cleaved by proteases during fusion. As mentioned above, CD9 and CD81 may also contribute to macrophage fusion. Additionally, DC-STAMP seems to be required for macrophage fusions. Mice lacking DC-STAMP are osteopetrotic and lack multinucleated osteoclasts and giant cells (Yagi et al. 2005). DC-STAMP is a seven-transmembrane receptor, somewhat similar to the HIV co-receptor CXCR4, but a ligand has yet to be identified.
These and many more molecules, including integrins, vacuolar ATPase and receptors and their ligands, as well as different signaling intermediates have been associated with cell fusions. With the exception of the syncytins, most of the mammalian molecules so far studied do not fulfill strict criteria for fusogens and several are dispensable for fusions (reviewed by Chen and Olson 2005; Chen et al. 2007; Oren-Suissa and Podbilewicz 2007). However, this may reflect molecular redundancy and does not definitely exclude that these molecules may participate in fusions. It is noteworthy that, in both man and mouse, two different syncytins are expressed and are both fusogenic. It is possible that, in the mammalian system, fusion requires a flotilla of molecules organized into membrane microdomains by proteins like CD9, encompassing receptors/ligands, signaling entities, proteases and fusogens of retroviral origin that are capable of forming alpha-helical bundles, which bring membranes closer. Syncytins and related retroviral envelope sequences may, in this connection, function both as fusogens and receptor ligands. However, it seems unlikely that as irreversible an event as a cell–cell fusion should depend upon a single receptor–ligand interaction. Thus, both facilitatory and inhibitory factors are expected to be part of the flotilla.
Cancer cell fusions
Studies by Mortensen et al. (2004) documented that human breast cancer cells fused with endothelial cells in culture. Stimulated by studies showing that syncytin-1 was involved in cytotrophoblast cell fusions (Blond et al. 2000; Mi et al. 2000), we examined whether a similar mechanism could account for cancer–endothelial cell fusions. Expression of syncytin-1 was documented in the breast cancer cell lines examined (MCF-7 and MDA-MB-231 cells; Bjerregaard et al. 2006, SK-BR-3 cells: Talts, Bjerregaard and Larsson: unpublished data). Moreover, we found that both tumor cells and endothelial cells expressed the syncytin-1 receptor ASCT-2. Use of phosphorthioate-protected syncytin-1 antisense oligonucleotides downmodulated syncytin-1 expression as measured by either quantitative RT-PCR or Western blotting and inhibited breast cancer–endothelial cell fusions, whereas a scrambled oligonucleotide control was without effect. Additionally, the syncytin-1 CHR peptide, referred to above, also inhibited the fusions whereas a control peptide was without effect (Bjerregaard et al. 2006). However, neither the antisense nor the CHR peptide experiments effected a total inhibition of cancer-endothelial cell fusions. There may be several reasons to this. Thus, the antisense oligonucleotide did not totally downmodulate syncytin-1 levels and could therefore not be expected to decrease cell fusions to zero and the lack of total inhibition by the CHR peptide could potentially be ascribed to proteolytic degradation. However, we also detected that the breast cancer cells produced syncytin-2 and cannot exclude that also this molecule contributed to the cancer–host cell fusions. Further experiments using shRNA-directed downmodulation of both syncytin-1 and -2 as well as of additional putative fusogenic retroviral sequence are now underway to test this. Nevertheless, these data provide strong evidence that syncytin-1 is involved in mediating cancer–endothelial cell fusions in vitro.
We next examined two series of human breast cancer patients for tumoral expression of syncytin-1 using a polyclonal antiserum raised to a synthetic nonapeptide derived from the syncytin-1 sequence. In addition, tumors were also screened for expression of ASCT-2 using a peptide antiserum. Preabsorption of the antisera with the corresponding peptides, but not with irrelevant peptides, abolished staining (Larsson et al. 2007b) (Fig. 6). The results showed that 38% of all breast cancer samples showed detectable staining for syncytin and that endothelial cells expressed ASCT-2 (Fig. 6). Moreover, significant expression of ASCT-2 was detected also in many tumor cells (Fig. 6). The degree of syncytin immunostaining was visually graded using coded specimens and statistical analysis showed that it correlated positively with disease-free survival of the patients (Larsson et al. 2007b). Multivariate analysis included age dichotomized at 40 years, tumor size dichotomized at 20 mm, grade and adjuvant therapy and identified syncytin expression as an independent prognostic indicator of increased disease-free survival Also when used as a continuous variable, syncytin expression emerged as a significant prognostic indicator for disease-free survival in the Cox model (P = 0.02) (Larsson et al. 2007b).
Fig. 6Human breast cancers immunocytochemically stained for syncytin-1 (a) or with antigen-preabsorbed antiserum (b) and for ASCT-2 (c, e) or with antigen-preabsorbed antiserum (d). Nuclei are lightly counterstained with haematoxylin. Note presence of syncytin immunoreactivity in tumor cells and of ASCT-2 immunoreactivity in endothelial cells (arrows) and in tumor cells
The involvement of syncytin-1 in tumor cell fusion events was subsequently confirmed by Strick et al. (2007), working with endometrial carcinomas. In their study, downmodulation of synctytin-1 expression also inhibited fusions between endometrial tumor cells. In agreement with findings on placental cells (vide supra), both cAMP elevating agents and estrogens upregulated syncytin-1 expression. However, only cAMP elevating agents stimulated cell fusions (Strick et al. 2007). This apparent discrepancy may possibly be ascribed to the fact that estrogen treatment upregulated TGFbeta, which interfered with syncytin-induced fusions. Thus, estrogen did, in fact, stimulate cell fusions if TGFbeta was immunoneutralized. Conversely, additions of TGFbeta 1 or 3 reduced fusions induced by cAMP-elevating agents. This effect was observed both in endometrial carcinoma cells and in trophoblast-derived cells. These results show that both cAMP and estrogens positively may regulate syncytin-1 expression in tumor cells and that the TGFbeta family may negatively regulate the fusogenic effects of syncytin-1 in both trophoblasts and cancer cells (Strick et al. 2007).
The role of syncytin-1 and cell fusions in cancer needs further study. Thus, our data show that syncytin-1 is not the only fusogenic protein expressed by breast cancer cells and the results presented by Strick et al. (2007) show that additional regulators, such as TGFbeta isoforms may be important modulators of cell fusions. So far, our breast cancer data indicate that syncytin-1 expression constitutes a positive prognostic factor. This is not the same as to say that cell fusions may be universally beneficial to cancers. First, syncytins may have additional effects within the tumor environment (Larsson et al. 2007a, b). Secondly, it seems likely that several factors within the cancer and its stroma (inactivated tumor suppressor genes, activated oncogenes, expression of fusogens and of CD9 and CD81 as well as TGFbeta) may act together to bring about a tumor profile that may be as diverse as the one demonstrated by the cell fusion experiments referred to above. Interestingly, however, expression of CD9 has been analyzed in a number of tumors and seems, like syncytin expression, to predict a good prognosis (Funakoshi et al. 2003; Hashida et al. 2002; Higashiyama et al. 1997; Houle et al. 2002; Miyake et al. 1995; Uchida et al. 1999). We propose that syncytins may be fusogens of importance to both trophoblast and cancer cell fusions and that they possibly also may mediate additional cell fusion events, acting in concert with other molecules with both enhancing and inhibitory regulatory effects.

cell fusion
cancer
syncytin
placenta
env-w


Tie-dyed1 and Sucrose export defective1 act independently to promote carbohydrate export from maize leaves

tie-dyed1 (tdy1) and sucrose export defective1 (sxd1) are recessive maize (Zea mays) mutants with nonclonal chlorotic leaf sectors that hyperaccumulate starch and soluble sugars. In addition, both mutants display similar growth-related defects such as reduced plant height and inflorescence development due to the retention of carbohydrates in leaves. As tdy1 and sxd1 are the only variegated leaf mutants known to accumulate carbohydrates in any plant, we investigated whether Tdy1 and Sxd1 function in the same pathway. Using aniline blue staining for callose and transmission electron microscopy to inspect plasmodesmatal ultrastructure, we determined that tdy1 does not have any physical blockage or alteration along the symplastic transport pathway as found in sxd1 mutants. To test whether the two genes function in the same genetic pathway, we constructed F2 families segregating both mutations. Double mutant plants showed an additive interaction for growth related phenotypes and soluble sugar accumulation, and expressed the leaf variegation pattern of both single mutants indicating that Tdy1 and Sxd1 act in separate genetic pathways. Although sxd1 mutants lack tocopherols, we determined that tdy1 mutants have wild-type tocopherol levels, indicating that Tdy1 does not function in the same biochemical pathway as Sxd1. From these and other data we conclude that Tdy1 and Sxd1 function independently to promote carbon export from leaves. Our genetic and cytological studies implicate Tdy1 functioning in veins, and a model discussing possible functions of TDY1 is presented.
Introduction
Sugars synthesized in photosynthetic leaf cells must be transported into the veins for distribution to nonphotosynthetic tissues. Maize (Zea mays) is a C4 plant which displays Kranz anatomy in its leaves (Esau 1977). Veins are surrounded by bundle sheath cells which in turn are surrounded by mesophyll cells. Mesophyll and bundle sheath cells cooperatively perform the reactions of photosynthesis and carbon assimilation. Sucrose is synthesized in the cytoplasm of mesophyll cells (Lunn and Furbank 1999), diffuses through plasmodesmata into bundle sheath cells, and then into vascular parenchyma cells (Russin et al. 1996). Maize is an apoplastic phloem loading species (Evert et al. 1978), whereby sucrose is exported from the vascular parenchyma cells to the apoplast by an unknown mechanism (see Lalonde et al. 2004 for discussion). Sucrose is then imported into phloem companion cells and/or sieve elements by sucrose transporters located in the plasma membrane (Aoki et al. 1999; Lalonde et al. 2004; Sauer 2007; Scofield et al. 2007). The energy to transport sucrose into the phloem is derived from H+-ATPases that generate the proton motive force across the plasma membrane (Bush 1993; Gaxiola et al. 2007). In addition, the abundance of the sucrose transporter that controls phloem loading is transcriptionally regulated by a sucrose-sensing signal transduction pathway (Chiou and Bush 1998; Vaughn et al. 2002; Ransom-Hodgkins et al. 2003). If carbon transport into the vein is blocked, carbohydrates accumulate in the photosynthetic cells, leading to down-regulation of photosynthetic gene expression and reduced chlorophyll levels (chlorosis) (Sheen 1990; Goldschmidt and Huber 1992; Riesmeier et al. 1994; Krapp and Stitt 1995; Burkle et al. 1998; Gottwald et al. 2000; Jeannette et al. 2000). Even though the transport pathway for assimilated carbon has been studied in maize leaves for almost 40 years (Hofstra and Nelson 1969), the mechanisms that regulate sucrose export from leaves remain poorly understood.
To date, two recessive maize mutants have been described with defects in carbon export from leaves. tie-dyed1 (tdy1) and sucrose export defective1 (sxd1) mutants develop nonclonal chlorotic leaf sectors that hyperaccumulate starch and soluble sugars (Russin et al. 1996; Provencher et al. 2001; Braun et al. 2006). Other mutants with variegated leaves have been characterized, but none have carbohydrate accumulation within the chlorotic regions (Rhoades and Carvalho 1944; Fisher and Eschrich 1985; Madore 1990; Yu et al. 2007). tdy1 and sxd1 both display growth-related defects such as reduced plant height and inflorescence development due to the retention of carbohydrates in leaf tissues. However, their phenotypes also show significant differences. sxd1 mutant leaves progressively exhibit chlorosis which initiates at the leaf tip and basipetally spreads toward the base. Interestingly, in sxd1 mutants, only the leaf minor veins are affected in phloem loading (Russin et al. 1996). The tdy1 mutant phenotype is distinct from sxd1 mutants in that large chlorotic regions develop throughout leaf blade tissue. Further, tdy1 chlorotic regions form only during a limited time as the leaf emerges from the whorl and, once formed, do not expand (Braun et al. 2006). Lastly, in tdy1 mutants, chlorotic tissues are often bounded by lateral veins, implicating this vein class in limiting the expansion of a tdy1 chlorotic sector (Baker and Braun 2007).
Sxd1 encodes tocopherol cyclase, an enzyme functioning in tocopherol (vitamin E) biosynthesis, and sxd1 mutants lack tocopherols (Sattler et al. 2003). Absence of tocopherols results in callose deposition over the plasmodesmata at the bundle sheath-vascular parenchyma cell interface of leaf minor veins (Russin et al. 1996; Botha et al. 2000; Provencher et al. 2001). This occludes these passageways and prevents sucrose from moving into the vein, resulting in the build-up of carbohydrates in photosynthetic cells. The function of tocopherols in preventing callose deposition in the phloem is evolutionarily conserved. RNAi-mediated suppression of tocopherol cyclase activity in potato results in callose deposition in vascular-associated cells and carbohydrate accumulation in the photosynthetic cells (Hofius et al. 2004). In addition, in Arabidopsis tocopherol cyclase mutants and other vitamin E deficient mutants, callose is deposited in phloem parenchyma transfer cells and the photosynthetic cells accumulate carbohydrates if the plants are subjected to nonfreezing low temperatures (Maeda et al. 2006). It is not known how lack of tocopherols causes callose deposition.
Because both mutants show defects in carbon export from leaves, we examined whether Tdy1 and Sxd1 function in the same pathway to limit leaf carbohydrate accumulation. Using aniline blue fluorescence microscopy and transmission electron microscopy (TEM) we determined that carbohydrate accumulation in tdy1 mutant leaves does not result from a similar impediment to phloem loading as in sxd1 mutants. To genetically test whether the two genes function in the same pathway, we analyzed F2 families segregating both mutations. We observed that the double mutant plants showed an additive interaction for growth-related phenotypes and soluble sugar accumulation indicating that the two genes function in separate genetic pathways. To determine if Tdy1 functions in the same biochemical pathway as Sxd1, we measured tocopherol levels in mutant plants. We found that tdy1 mutants contain wild-type levels of tocopherols, indicating that Tdy1 does not function in the same biochemical process as Sxd1. Thus, from the combination of cellular, genetic and biochemical investigations we conclude that Tdy1 and Sxd1 function independently to promote carbon export from leaves. Additionally, genetic and cytological studies suggest that Tdy1 functions within the veins, and a model of possible TDY1 functions is discussed.
Materials and methods
Plant materials
Plants were grown in the summer in the Rock Springs Agronomy Farm, Pennsylvania State University, and in the winter in Juana Diaz, Puerto Rico. The tdy1-Reference (tdy1-R) (Braun et al. 2006) and sxd1-1 (Russin et al. 1996) mutations were backcrossed to the B73 inbred line five times prior to crossing together. F2 families were generated by crossing tdy1-R and sxd1-1 mutant plants and self-pollinating the F1 individuals.
Morphological analyses
Plant height was measured from the soil surface to the tip of the central spike of the tassel. Tassel length was measured from the node of the flag leaf to the tip of the central spike. Anthesis was recorded as the day of first pollen shed. Plants were scored as having produced ears if either (1) ears were clearly visible, (2) silks were visible emerging from the leaf sheath, or (3) ears developed such that they were identifiable by peeling back the ear leaf sheath. For wild-type and tdy1-R single mutants n = 18 for all measurements. For sxd1-1 single mutants, 12 of the 16 plants produced a tassel that shed pollen, and 13 produced an ear. For tdy1-R; sxd1-1 double mutants, 7 of the 15 plants shed pollen and 8 made an ear. Barren tassels that did not shed pollen were not included in the tassel length or anthesis measurements.
Aniline blue staining
Leaf strips 1 cm in width were cut from mutant and wild-type leaves. The abaxial epidermis and the mesophyll cells directly beneath it were gently scraped away with a single-edge razor to expose the inner leaf tissues. The leaf strip was placed on a slide, briefly stained with a 0.05% aniline blue and immediately viewed using a 360–370-nm excitation filter and a 420-nm long pass emission filter on a Nikon Eclipse 80i fluorescent microscope. Epifluorescent illumination was provided by a 100-W mercury lamp, and images were recorded using a DXM1200F Nikon digital camera.
TEM analyses
For tissue fixation, leaf samples were diced into 3 × 1 mm pieces, placed in 4% glutaraldehyde, 1% paraformaldehyde, 0.3% Tween 20, 50 mM sodium cacodylate, pH 7.4, and vacuum infiltrated on ice for 4–6 h. Samples were post-fixed in 1–2% osmium tetroxide at 4°C overnight, subsequently dehydrated through a graded acetone series and embedded in Spurr’s epoxy resin (Spurr 1969). Thin sections (90–100 nm) were cut on a Leica Ultracut E ultramicrotome with a glass knife and lifted onto 200-mesh copper grids. The grids were stained in 2% uranyl acetate, followed by Reynold’s lead citrate and observed with a JEOL JEM 1200 EXII at an accelerating voltage of 80 kV.
Chlorophyll quantification
Relative levels of total chlorophylls were quantified from leaves using a SPAD 502 Data Logger Chlorophyll Meter (Spectrum Technologies). For each tissue, 30 samples were measured, and the experiment was performed three times. Representative data from one replicate are shown.
Sugars and starch quantification
Leaf tissues were collected at the end of the photoperiod, weighed and frozen at −80°C. Carbohydrate extraction was performed according to Dinges et al. (2001). Sugars and starch were quantified using commercial assay kits according to the manufacturer’s instructions (R-Biopharm). Six samples from each tissue type were measured in triplicate.
For visualizing starch in leaves, samples were collected at the end the photoperiod, decolored by boiling in 95% ethanol and stained with iodine-potassium iodide (IKI) (Ruzin 1999).
Tocopherol quantification
Leaf discs of 11 mm were harvested with a cork borer from leaf tissues, weighed and immediately frozen on dry ice. Total lipids were extracted from collected tissues and tocopherols measured according to (Sattler et al. 2003).
Results
tdy1-R and sxd1-1 single mutants both display variegated chlorotic leaves
tdy1-R and sxd1-1 mutants show striking parallels in their leaf phenotypes. Both develop chlorotic sectors that hyperaccumulate starch and soluble sugars in leaf blades (Fig. 1). However, there are also distinctions. Whereas wild-type leaves have a uniform, dark green color, sxd1-1 mutant leaves develop a chlorotic phenotype that shows a continuum of expression from strongest at the tip to mildest toward the base (Fig. 1a, b). In addition, at the tip and leaf margins, the chlorotic tissues progressively accumulate anthocyanin and excess starch (Fig. 1b, e). In sxd1-1 leaves, no clear, delineated boundaries occur between green and chlorotic regions. Conversely, tdy1-R mutants display a variegated pattern of chlorotic and normal appearing green regions throughout leaf blade tissue (Fig. 1c). Additionally, the boundaries between tdy1-R chlorotic and normal appearing green tissue are sharp and distinct, and they tend to occur at lateral veins (Baker and Braun 2007). As in sxd1-1 mutants, chlorotic regions in tdy1-R mutants hyperaccumulate starch relative to green tissues (Fig. 1f). tdy1 and sxd1 are the only known leaf variegation mutants that hyperaccumulate carbohydrates in any plant; therefore, we investigated whether Tdy1 and Sxd1 function in a common pathway influencing carbon partitioning in leaves.
Fig. 1tdy1-R and sxd1-1 leaves display chlorotic sectors that hyperaccumulate carbohydrates. a Wild-type leaf showing uniform dark green color. b sxd1-1 mutant leaf displaying a chlorotic gradient, strongest at the tip to mildest toward the leaf base. Anthocyanins accumulate in the chlorotic tissues at the leaf tip and margins. c tdy1-R mutant leaf containing variegated chlorotic and green sectors throughout the leaf blade. d–f Cleared, IKI stained leaf tissues showing that mutant chlorotic regions hyperaccumulate starch. d Wild type. e sxd1-1. f tdy1-R. Scale bars represent 4 cm
Carbohydrate accumulation in tdy1-R mutants occurs by a different mechanism than in sxd1-1 plants
To address whether the two genes function in the same pathway, we investigated if the mechanism for carbohydrate accumulation was similar in the two mutants. Carbohydrate accumulation in sxd1-1 mutants results from a blockage in the symplastic pathway of sucrose movement (Russin et al. 1996; Botha et al. 2000; Provencher et al. 2001). Specifically, at the bundle sheath–vascular parenchyma cell interface of leaf minor veins, callose is deposited over the plasmodesmata in bundle sheath cells. This blockage prevents sucrose from being loaded into the phloem and results in carbohydrate accumulation in the photosynthetic cells. To determine if the tdy1-R leaf phenotype is similarly caused by callose deposits plugging the plasmodesmata, we performed aniline blue staining and fluorescence microscopy. Aniline blue binds callose and fluoresces a blue–white color under UV light. In wild-type plants, only the sieve plates in the end walls of phloem sieve elements stain positively for callose (Fig. 2b). As previously reported, sxd1-1 mutants display many punctate callose deposits at the vascular parenchyma-bundle sheath cell interface of minor veins (Fig. 2c) (Botha et al. 2000). In tdy1-R chlorotic regions, no ectopic callose deposits were detected at this interface (Fig. 2d). Further, no callose deposits were observed in any cells of minor or lateral veins in tdy1-R chlorotic tissues, nor in any cells of tdy1-R green regions (data not shown). These data suggest that tdy1-R mutants do not accumulate carbohydrates due to callose blocking symplastic sucrose movement.
Fig. 2Aniline blue staining of ectopic callose deposits in leaf minor veins. a Cross section of a wild-type minor vein shown by UV autofluorescence indicating the cell types and orientation of view (arrow) for b–e. Abaxial cells were removed below the black dotted line to view the bundle sheath–vascular parenchyma cell interface. X xylem, P phloem, VP vascular parenchyma, BS bundle sheath, M mesophyll. b–eAniline blue fluorescence images of paradermal sections along minor veins. b Wild-type cells lack punctate callose deposits indicating no blockages along the symplastic pathway. The bright fluorescence observed in the phloem corresponds to sieve plates (arrowhead). c sxd1-1 minor veins contain many callose deposits at the bundle sheath–vascular parenchyma cell interface (arrow). d tdy1-R chlorotic tissue lacks callose deposition over plasmodesmata in any cells. e tdy1-R; sxd1-1 double mutants contain callose deposits (arrow) at the bundle sheath–vascular parenchyma cell interface comparable to the level seen in sxd1-1 single mutants. Scale bar in a represents 50 μm, and in b–e 10 μm
Although we did not observe ectopic callose deposition in any cells in tdy1-R mutant leaves using aniline blue staining, it is possible that symplastic sucrose transport is precluded due to a different defect in plasmodesmatal structure. To investigate this possibility, we performed TEM to inspect the plasmodesmata ultrastructure along the symplastic pathway. In minor veins from wild-type leaves, the plasmodesmata between bundle sheath and vascular parenchyma cells were unobstructed, spanned the cell wall and connected the cytoplasms between the two cells (Fig. 3a). Consistent with previous reports, minor veins from leaf tips expressing the sxd1-1 mutant phenotype showed occlusions over the plasmodesmata on the bundle sheath cell side of the bundle sheath–vascular parenchyma cell interface (Fig. 3b) (Russin et al. 1996; Botha et al. 2000; Provencher et al. 2001). In tdy1-R mutant leaves, we found no occlusions over the plasmodesmata or alterations in their structure between the bundle sheath and vascular parenchyma cells in minor veins (Fig. 3c). In examining plasmodesmata along the symplastic pathway at all cellular interfaces in tdy1-R leaves, we did not observe any structural perturbations or occlusions (Fig. 3e–j). These data indicate that carbohydrate retention in tdy1-R mutant leaves does not appear to result from physical blockage or plasmodesmatal structural changes. Together, the aniline blue staining and TEM studies indicate that the cellular basis for the tdy1-R and sxd1-1 phenotypes is different.
Fig. 3TEM images of cellular interfaces along the symplastic pathway of minor veins. a-d Bundle sheath–vascular parenchyma cells. Arrows indicate the location of the plasma membrane in the bundle sheath cell. a Wild-type plasmodesmata span the cell wall, lack occlusions, and connect with the plasma membrane. b sxd1-1 chlorotic leaf tips contain occlusions over the plasmodesmata on the bundle sheath cell side of the cell wall. c tdy1-R chlorotic sectors contain normal appearing plasmodesmata that lack occlusions. Note the plasmodesmata span the cell wall and connect with the plasma membrane. d tdy1-R; sxd1-1double mutant chlorotic tissues contain deposits over the plasmodesmata in the bundle sheath cell similar to sxd1-1 single mutants. Additional cellular interfaces along the symplastic pathway of wild type (e, g, i) and tdy1-R chlorotic tissues (f, h, j) are shown. In e–j the plasmodesmata spanned the cell wall connecting the cytoplasms of adjacent cells and lacked occlusions. e, f BS–BS cells, g, h BS–M cells, i, j M–M cells. BS bundle sheath, M mesophyll. Scale bars represent 200 nm
Double mutant plants show more severe growth defects than either single mutant
Both tdy1-R and sxd1-1 single mutant plants have growth-related defects due to the retention of carbohydrates in leaves. To determine whether the two genes function in the same genetic pathway promoting carbon export from leaves, we created F2 families segregating both mutations. Both mutations are recessive and the F2 segregation ratio fits the expected 9:3:3:1 prediction, indicating that neither mutant shows epistasis (Table 1). sxd1-1 is a molecular null allele lacking transcripts (Provencher et al. 2001), and based on dosage analysis, tdy1-R is a genetic null allele (Braun et al. 2006). If Tdy1 and Sxd1 act in a linear genetic pathway, we would expect that the phenotype of the double mutant would be similar to one or the other mutant. If the two genes function in separate pathways, then we predict that the double mutant would be more severe than either single mutant.
Table 1Genetic segregation in F2 familiesFamily #TotalWild typetdy1-Rsxd1-1tdy1-R; sxd1-1χ2DB 55039348221672.0P > 0.05DB 55129552231552.1P > 0.05DB 55239355181550.6P > 0.05Total2811556346172.9P > 0.05χ2 analyses of tdy1-R and sxd1-1 segregation in three F2 families conform to a 9:3:3:1 expectation. Observed numbers of individuals are shown
To examine the interactions between tdy1-R and sxd1-1, we quantified the growth parameters of plant height, tassel size, time to anthesis and ear production (Fig. 4, Table 2). At the whole plant level, the double mutants express a more severe phenotype and show a stronger reduction in growth characteristics than either single mutant (Fig. 4a). Compared to wild-type, tdy1-R single mutants have an 11% decrease, sxd1-1 mutants have a 24% reduction, and the tdy1-R; sxd1-1 double mutant plants show the greatest reduction in plant height of 35.5%. Similar to plant height, tassel size is reduced 13% in tdy1-R plants, 29% in sxd1-1 mutants, and 50% in tdy1-R; sxd1-1 double mutants (Fig. 4c, Table 2). In addition, approximately half of the tdy1-R; sxd1-1 double mutant plants produced very reduced tassel branches lacking spikelets (Fig. 4d). This phenotype was observed in 25% of the sxd1-1 single mutants. Correlated with the tassel height defects, both mutations retarded time to flowering. Both tdy1-R and sxd1-1 single mutants first shed pollen eight days later than wild-type siblings (Table 2). Double mutant plants were delayed even further by approximately another 5 days. Similarly, wild-type and tdy1-R mutant plants invariably made ears, and sxd1-1 mutants produced ears 81% of the time. In contrast, only 53% of the double mutant plants produced an ear (Table 2). All of the growth defects observed in the single and double mutants are consistent with a restriction in assimilates transported to the growing portions of the plants. The double mutant plants showed the most severe phenotypes suggesting they have a greater reduction in carbohydrates exported out of leaves than in either single mutant. In support of this, the leaves of tdy1-R; sxd1-1 double mutant plants are more strongly chlorotic than either single mutant, with a corresponding reduction in the amount of green tissue (compare Figs. 4b with 1b, c). In double mutant leaves, the tip to base gradient of chlorosis and anthocyanin pigmentation characteristic of the sxd1-1 phenotype is superimposed on the tdy1-R chlorotic and green sectored pattern. Double mutant leaves still contain some green tissues located at the leaf base as in sxd1-1 single mutants; however, the boundaries between the chlorotic and green regions are sharp and distinct as in tdy1-R mutants (Fig. 4b). Thus, the double mutant leaves express the phenotypes of both single mutants, and double mutant plants display stronger growth retardation than either single mutant.
Fig. 4Phenotypic comparisons among plants from a F2 family segregating tdy1-R and sxd1-1 mutants. In a and c the order from left to right corresponds to wild type, tdy1-R, sxd1-1, and tdy1-R; sxd1-1 double mutant. a Double mutants show the greatest reduction in plant height compared with either single mutants or wild-type siblings. b Double mutant leaves are severely chlorotic with lighter green tissues restricted to the leaf base. Distinct borders between the chlorotic and green tissues characteristic of tdy1-R are visible at the leaf base. The strong tip to base chlorosis gradient of sxd1-1 expression is evident. c Tassel height is most strongly reduced in tdy1-R; sxd1-1 double mutants compared with sxd1-1 and tdy1-R single mutants or wild-type siblings. d Approximately half of the tdy1-R; sxd1-1 double mutant plants produced barren tassels lacking spikelets. Scale bars represent 4 cmTable 2Morphometric analyses in F2 familiesPhenotypePlant heightwt%Tassel lengthwt% AnthesisEars (%)Wild type220.4a ± 0.3910055.2a ± 0.1410071.9a ± 0.06 100tdy1-R196.8b ± 0.9289.247.9b ± 0.28 86.880.3b ± 0.13 100sxd1-1168.9c ± 1.2476.639.3c ± 0.1871.280.3b ± 0.0981tdy1-R; sxd1-1142.2d ± 0.7964.526.8d ± 0.34 48.684.9c ± 0.1353Values are the means ± SE. Plant height and tassel length are measured in cm. Anthesis is measured as the number of days after planting until first pollen shed. Percent ears indicates the frequency each phenotypic class produced an ear. Different letters in superscript within a column denote statistical significance determined using Student’s t test
Since double mutants have more extensive chlorosis in their leaves, we ascertained whether the callose deposits at the vascular parenchyma-bundle sheath cell interface observed in sxd1-1 single mutants were more severe in the double mutants. Using aniline blue fluorescence, the double mutants showed no enhancement of the callose deposition phenotype and appeared similar to sxd1-1 single mutants (Fig. 2c, e). Similarly using TEM, we found blockages over the plasmodesmata on the bundle sheath cell side of the bundle sheath–vascular parenchyma cell interface of minor veins in the double mutants comparable to what we observed in sxd1-1 single mutant leaves (Fig. 3b, d). In neither case was there any increase in the severity of the phenotype. Hence, tdy1-R and sxd1-1 do not display a synergistic interaction as the double mutant leaves express both single mutant phenotypes. Rather, these data suggest that the phenotypes observed in the double mutants are additive, and that the genes act in independent pathways.
Leaf tips of double mutant plants have the greatest reduction in chlorophyll levels
tdy1-R and sxd1-1 single mutants both show variegated chlorotic regions in their leaves (Fig. 1b, c). We quantified total chlorophyll levels to determine whether the double mutants were more strongly affected than either single mutant (Table 3). We observed a decreasing series in chlorophyll content with the strongest reduction in the tips of double mutant leaves < the chlorotic regions of either tdy1-R or sxd1-1 single mutant leaves < the green base of sxd1-1 or double mutant leaves < green regions of tdy1-R mutants or wild-type leaves. The leaf bases of sxd1-1 mutants and the double mutants displayed a mild chlorosis phenotype with approximately 70% as much chlorophyll as wild-type leaves indicating that these tissues are affected by the sxd1-1 mutation. As the tips of double mutant leaves showed a more severe chlorosis phenotype than either single mutant, it suggests that Tdy1 and Sxd1 function in separate pathways.
Table 3Chlorophyll quantification in F2 plantsPhenotypeChlorophyllWild type55.7a ± 0.55tdy1-R green54.8a ± 0.57sxd1-1 green41.0b ± 0.75tdy1-R; sxd1-1 green42.2b ± 0.67tdy1-R chlorotic20.9c ± 0.62sxd1-1 chlorotic21.5c ± 0.66tdy1-R; sxd1-1 chlorotic11.4d ± 0.64Chlorophyll levels were measured in different phenotypic (green or chlorotic) tissues from different genotypes. Values represent the means ± SE in relative units (n = 30). Statistical significance was determined using Student’s t test and indicated by different letters in superscript
Sugar accumulation in double mutant leaves is greater than in either single mutant
Both single mutants accumulate excess starch in the chlorotic regions of their leaves (Fig. 1e, f). To investigate whether Tdy1 and Sxd1 act in the same or separate genetic pathways limiting carbohydrate accumulation in leaves, we determined the starch and soluble sugars content in different phenotypic regions of F2 plants (Fig. 5; Table S1 in electronic supplementary material). For both the green regions and the chlorotic regions, we found that the double mutants contained similar amounts of starch as the single mutants. In contrast, we determined that the levels of soluble sugars differed with respect to genotype and phenotype. As previously reported, we found green regions of tdy1-R leaves had sucrose, glucose and fructose levels indistinguishable from wild type (Braun et al. 2006). However, green leaf bases of the double mutants and sxd1-1 single mutants contained approximately fourfold higher levels of glucose and fructose as wild type or tdy1-R green tissues. Among the chlorotic leaf tissues, the double mutant leaf tips had the highest concentrations of sugars, with approximately 33% greater amounts of glucose, and a significant increase in sucrose content compared with either single mutant. Double mutant leaf tips also contained ∼30% more fructose than sxd1-1 leaf tips and approximately two-fold greater levels than in tdy1-R chlorotic regions. Overall, sugar concentrations in green tissues of double mutant leaves are more similar to sxd1-1 leaf bases than to tdy1-R green regions, whereas in chlorotic tissues, the double mutants contain more soluble sugars than either single mutant. Because the double mutants show greater individual sugar accumulation than either single mutant, these data support the hypothesis that Tdy1 and Sxd1 function in independent pathways promoting carbon export from leaves.
Fig. 5Carbohydrate quantification in leaves of wild-type, tdy1-R, sxd1-1 and double mutant plants. Units for all panels are mg carbohydrate/g fresh weight. Values are the means of 18 samples ± SE. Abbreviations are wt, wild-type, dm, tdy1-R; sxd1-1 double mutant. Different letters indicate statistically significant differences between samples determined using Student’s t test
Tdy1 does not function in the same biochemical pathway as Sxd1
Sxd1 encodes tocopherol cyclase, the penultimate enzyme in the vitamin E biosynthesis pathway, and sxd1 mutants lack tocopherols (Sattler et al. 2003). To determine whether Tdy1 functions in the same biochemical pathway as Sxd1, we quantified tocopherol levels in tdy1-R plants (Table 4). We found no statistically significant changes in tocopherol quantities in tdy1-R chlorotic or green regions in comparison to wild-type tissue. sxd1 mutants had undetectable levels of tocopherols as previously reported (Sattler et al. 2003). Hence, tdy1-R does not have a lesion in tocopherol synthesis or accumulation, and Tdy1 does not function in the same biochemical pathway as Sxd1.
Table 4Tocopherol quantificationPhenotypeTocopherolWild type118.6a ± 3.59tdy1-R green101.0a ± 3.17tdy1-R chlorotic112.3a ± 6.17sxd1-1ndValues are means of six samples ± SE measured in μg/g fresh weight. nd indicates none detected. Values indicated by letter “a” in superscript were not statistically different as determined using Student’s t test
Discussion
Tdy1 and Sxd1 independently promote carbon export in maize leaves
tdy1 and sxd1 are the only known mutations in plants that confer a variegated leaf phenotype with sectors that hyperaccumulate carbohydrates. Using a combination of biochemical, cytological and genetic analyses we investigated whether Tdy1 and Sxd1 act in a common pathway. Whereas sxd1 mutants lack tocopherol, we determined that tdy1-R mutants accumulate wild-type levels of tocopherols. Therefore, Tdy1 does not function in the same biochemical process as Sxd1. sxd1 mutants accumulate carbohydrates in leaf tissues due to callose being deposited over the interface of bundle sheath-vascular parenchyma cells in leaf minor veins which blocks sucrose symplastic movement. Using aniline blue fluorescence microscopy we did not identify any ectopic callose deposition in tdy1-R mutant leaf tissues. Moreover, using TEM we did not observe any structural alterations to the plasmodesmata, indicating that the symplastic loading pathway is not physically impeded. This suggests that tdy1-R mutants do not accumulate carbohydrates due to an occlusion, but rather due to a different cellular mechanism. This was confirmed by double mutant analyses which showed that the two mutations had an additive interaction. We found that the double mutants expressed both single mutant leaf phenotypes, that the reductions in plant height, tassel size and chlorophyll levels were additive, and that the accumulation of sugars in chlorotic leaf tissues of the double mutants was greater than in either single mutant. In addition, half of the double mutants produced ears whereas the tdy1-R mutant frequency was indistinguishable from wild type and 81% of the sxd1-1 single mutants made ears. The increased frequencies in the failure to make ears and production of barren tassels observed in the double mutants are most likely explained by an additive retention of carbohydrates in leaf tissues and a concomitant failure to export sucrose to the developing inflorescences. Collectively, these data lead us to conclude that Tdy1 and Sxd1 function independently, and that tdy1 defines a distinct genetic pathway affecting carbon partitioning in maize leaves.
Chlorotic leaf tissues do not retain sink identity
We previously suggested that one possibility to explain the excess carbon accumulation in tdy1-R chlorotic leaf sectors was a block in the developmental transition from sink to source identity, such that the chlorotic sectors continued to import photoassimilates from neighboring green tissues (Braun et al. 2006). If the chlorotic tissues were to remain sinks, the large amounts of green source tissues could potentially provide the carbohydrates accumulating in the chlorotic regions. However, based on our genetic analyses, our data do not support this hypothesis. In the tdy1-R; sxd1-1 double mutants, the amount of green leaf tissue is greatly reduced and is restricted to a narrow region at the very base of the leaves. Additionally, this green tissue displays mild chlorosis and an increase in carbohydrate accumulation indicating that it is perturbed in carbon export. As this tissue is proximal to and up to a meter distant from the chlorotic tip of adult leaves, it is highly unlikely to be providing the carbohydrates that accumulate in the cells near the tip. Furthermore, half of the double mutant plants produced a tassel and an ear. These are sink tissues dependent on the photosynthetic source leaves for their carbohydrates. Because the great majority of the area of double mutant leaves is severely chlorotic, yet still must be capable of exporting photoassimilates to produce the developing reproductive tissues, our data argues against the chlorotic tissues remaining sinks and does not support the hypothesis that Tdy1 functions to regulate the sink to source transition. Rather, the new data lead us to favor the hypothesis that the cells in the tdy1 chlorotic region are source tissues that are partly blocked in carbon export capacity.
A model for TDY1 function
In wild-type maize leaves, sucrose is synthesized in mesophyll cells, diffuses through plasmodesmata into bundle sheath cells and then into the vascular parenchyma cells (Fig. 6a). Sucrose is exported from the vascular parenchyma cell to the apoplast, and then loaded into the phloem by sucrose transporters in the plasma membrane of the companion cell and/or sieve element (Lalonde et al. 2004; Sauer 2007). sxd1 mutant leaf tips are blocked in this pathway at the bundle sheath–vascular parenchyma cell interface (Fig. 6b) (Russin et al. 1996; Botha et al. 2000; Provencher et al. 2001).
Fig. 6Model for TDY1 functions in phloem loading of sucrose. Diagrams represent the pathway of sucrose movement from photosynthetic cells into the phloem. M mesophyll, BS bundle sheath, VP vascular parenchyma, CC companion cell, SE sieve element. Green and yellow ovals in M and BS cells represent chloroplasts. White circles within chloroplasts represent starch grains. Magenta box on VP plasma membrane represents sucrose efflux transporter, and blue boxes on CC and SE plasma membranes depict sucrose transporters. Arrows indicate the direction of sucrose movement. a Wild-type tissue with normal, unimpeded sucrose movement into the phloem. b sxd1-1 mutant chlorotic tissue has callose deposits (red box) over the plasmodesmata at the BS–VP cell interface blocking sucrose movement and resulting in carbohydrate accumulation in M and BS cells. c tdy1-R chlorotic tissue has normal appearing plasmodesmata and lacks callose deposits. We hypothesize that either the sucrose efflux transport step is inhibited (red X) or the CC–SE sucrose transporters (orange X) fail to load sucrose into the phloem. This would result in the carbon hyperaccumulation observed in the photosynthetic cells
tdy1 chlorotic sectors accumulate excess carbohydrates indicating that they have a defect in phloem loading. In agreement with this, we previously localized the site of TDY1 function to the innermost tissue layer of leaves comprised of the veins, bundle sheath cells, and interveinal mesophyll cells (Baker and Braun 2007). Due to the limits of the experiment it was not possible to further delineate where within this tissue layer TDY1 acts. However, the cytological investigations presented resolve this ambiguity. As we observed no blockages or alterations to plasmodesmatal structure along the sucrose symplastic pathway in tdy1-R leaves, the carbon accumulation points to a defect at a later step in phloem loading. Nonetheless, even though the plasmodesmata appear unobstructed, we cannot be certain that they are functional. However, we propose that a molecular blockage of plasmodesmata precluding sucrose movement is an unlikely scenario for the following reason. In sxd1 mutants, specific plasmodesmata are occluded and unable to transit sucrose, causing the vascular parenchyma cells to appear plasmolysed due to their low osmotic concentration (Russin et al. 1996). Accordingly, we carefully inspected for, but never observed, plasmolysis of any vascular parenchyma, mesophyll or bundle sheath cells in tdy1 mutants in our microscopic analyses. Therefore, these data suggest that sucrose is able to pass freely between cells along the symplastic pathway, and hence that TDY1 likely functions in the veins (Fig. 6c).
From the variegated phenotype of tdy1 mutants, we hypothesize that TDY1 acts as a regulator of sucrose transport rather than as a transporter itself, as loss of transporter function leads to uniform carbohydrate accumulation and chlorosis phenotypes (Riesmeier et al. 1994; Burkle et al. 1998; Gottwald et al. 2000; see Braun et al. 2006; Baker and Braun 2007 for discussion of mechanisms for tdy1 variegation). Several possibilities are envisioned for Tdy1 function. The first possibility is that Tdy1 may regulate sucrose efflux out of vascular parenchyma cells. Failure to export sucrose to the apoplast would create a bottleneck restriction leading to the build-up of sucrose in the vascular parenchyma cells and, in turn, to carbohydrate accumulation in the bundle sheath and mesophyll cells. The second possibility is that Tdy1 may function to regulate the activity of the H+-ATPases or sucrose transporters within the companion cells and sieve elements. Lack of function of either the H+-ATPases or sucrose transporters results in failure to load sucrose into the phloem and thereby to excess accumulation in the apoplast. Sucrose accumulation in the apoplast would shift the equilibrium for sucrose export from vascular parenchyma cells leading to increased concentration within these cells, and ultimately to carbon accumulation in the photosynthetic cells. In support of this, antisense expression or mutation of a phloem H+-ATPase or sucrose transporters causes an accumulation of carbohydrates in photosynthetic cells and chlorosis (Riesmeier et al. 1994; Burkle et al. 1998; Gottwald et al. 2000; Zhao et al. 2000). A third possibility is that TDY1 may function in the sucrose signal transduction pathway that regulates the abundance of the sucrose transporter controlling phloem loading (Chiou and Bush 1998; Vaughn et al. 2002; Ransom-Hodgkins et al. 2003).
Our analyses of the tdy1-R mutant phenotype and genetic interactions between tdy1-R and sxd1-1 have shown that Tdy1 acts independently of Sxd1. Thus, the tdy1 mutation identifies a separate genetic pathway limiting carbon accumulation in maize leaves. Future work characterizing the molecular function of Tdy1 will determine the mechanism by which Tdy1 influences carbon partitioning. Understanding the control of carbon export from leaves will lead to novel strategies to manipulate carbon allocation and biomass deposition which may have applications in the production of biofuels.
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tie-dyed1
sucrose export defective1
maize
carbohydrate accumulation
carbon partitioning


Effect of sesamin on serum cholesterol and triglycerides levels in LDL receptor-deficient mice

Background Sesamin, a major lignan from sesame seeds has been associated with cholesterol reduction in previous reports, but recent studies suggested differences in the response to sesamin intake depending on the model studied as well as the nature of the sesamin preparation used.
Introduction
The research on natural substances affecting cholesterol metabolism for prevention of hypercholesterolemic atherosclerosis has particular therapeutic importance. In particular, the identification of dietary components that can be added to foods to lower or regulate cholesterol levels has gained special interest.
Sesame seeds in general and its major lignan sesamin in particular, have been associated with various biochemical actions, mainly related to lipid metabolism, including a hypocholesterolemic effect in both human and animal-based trials (see [1, 2] for review). Although the general hypolipidemic effect of dietary sesamin seems to be clear in studies with rats, recent studies pointed out discrepancies in the response to sesamin between animal species [3] raising doubts about a possible extrapolation of these results to humans. Additionally, the heterogeneous nature of the sesamin preparations used in the different studies seems to be another point to consider, since different biological actions and mechanisms have been proposed for the two epimers of dietary sesamin [4]. In this study, we aimed to further investigate the hypocholesterolemic action of sesamin minimizing the above described variables and using only pure sesamin epimer and selecting low density lipoprotein (LDL) receptor-deficient mice as experimental animals. This model aims to reproduce homozygous familial hypercholesterolemia and it is characterized by decreased removal of cholesterol from the circulation due the lack of functional LDL receptors [5] that leads to increased serum cholesterol levels.
Materials and methods
Animal model
Female ldlr−/− mice (strain number, B6.129S7-Ldlrtm1Her) quality SPF were obtained from the Jackson Laboratory (Bar Harbor, Maine, US), and maintained at the Central Animal Laboratory of Raisio Benecol Ltd. (Raisio, Finland) in accordance with European guidelines (European Treaty Series No. 123, EU No 609/86, Official Journal of the European Communities No. L 358) approved by the Animal Care and Use Committee of the University of Turku (Finland) approval number 1263/02. Animals were 17–18 weeks at the beginning of the study/15–20 g and after 4 weeks of acclimatization, mice were housed (Polycarbonate Macrolon III, Scanbur AS, Denmark) in groups of 1–2/cage and randomly given free access to I–IV diets for 4 weeks. Ambient temperature was 21 ± 3°C, humidity ranged 55 ± 15% and illumination consisted in 12-h dark/light cycle. Free access to community tap water was allowed, except during the experiments. Blood samples were collected for the determination of the concentrations of serum cholesterol and triglycerides at the beginning and at the end of the 4-week dietary intervention. Mice were sedated with CO2/O2-mixture and the blood samples were collected from the orbital sinus (at the beginning) or through a cardiac puncture (at the end). At the beginning of the study the blood was collected into 500 µl gel tubes and allowed to clot at least for 1 h. Samples were then centrifuged for 15 min at 4000 rpm. Serum was separated and stored at −20°C until analyses. At the end of the intervention, blood samples were collected in heparin tubes and centrifuged within 1 h. The plasma samples were frozen at −20°C and stored until analyses. About 50 µl was used for cholesterol and triglyceride analyses and the rest was stored at −20°C for further analyses. During the intervention, the animals were weighed twice a week and observed daily (morning and afternoon) for general well being.
Test compounds and diets
Plant-derived stanol ester was produced by Raisio Benecol Ltd. (Raisio, Finland). Sesamin epimer (CAS No. 607–80-7, >95% purity, Fig. 1A) was synthesized by Industrial Research Ltd. (Wellingtown, New Zealand). During acclimatization/isolation period the diet was RM1 (E) SQC, (Special Diet Service, Witham Essex, UK). Experimental atherogenic diets containing 0.25% of cholesterol were prepared at Raisio Benecol Ltd. based on Clinton-Cybulsky diet premix (D12106pxc, Research Diets Inc., New Brunswick, NJ) in which the test compounds were administered, as follows: control (I) diet (n = 8), stanol ester (II) diet (n = 8, 7 mg/kg), sesamin (III) diet (n = 8, 1 mg/kg) and combined (IV) diet (n = 7, 7 mg/kg stanol ester and 1 mg/kg sesamin). Diets were stored at +2 to +8°C and allowed to warm to room temperature before delivery to the animals (twice a week) in 300-ml glass cups attached on the cage floor.
Fig. 1Chemical structure of sesamin epimers (A) sesamin and (B) episesamin (or asarinin)
Rationale for dose selection
Fat and cholesterol levels in the atherogenic diet were based on the literature [6]. The use of stanol ester as control test model has been proven several times to induce cholesterol reduction (Raisio Benecol Ltd., unpublished results) and the concentration in this study was selected to exert sufficient but not the maximal response, enabling the possible combination effect with sesamin. Sesamin concentration, that was adopted from previous reports [7–9] allowing comparison of the results was also considered as a rationale level to be used in clinical trials.
Analysis
Serum cholesterol and triglycerides were determined by Ecoline CHOD-PAP and Ecoline 25 GPO-PAP assay kits (1.14856.0001, Merck KGaA, Darmstadt, Germany), respectively. Control serum was Qualitrol HS N (Merck KGaA, Darmstadt, Germany). Serum metabolic profile of lignans was carried out by HPLC with coulometric electrode array detection [10] and the absence of plant lignans other than sesamin epimer in the diets was confirmed by isotope-dilution GC-MS [11].
Statistics
Graph Pad Prism software version 3.02 (GraphPad Software Inc., CA) was used for statistical data treatment. The differences in changes from the basal concentrations during the intervention between the groups were analyzed by ANOVA. Pair-wise comparisons between the groups were performed by Tukey’s Multiple Comparison Test. Two-way ANOVA was used to analyze differences in the body weights between the groups. A P-value less than 0.05 was considered as statistically significant. All analyses were performed as two-sided tests.
Results
Body weight gain
The body weights of the animals increased in all groups during the 4-week intervention period (Fig. 2). The increment tendency in group I was less than those in the other groups but analysis did not reveal a significant group × week interaction (P = 1.00). However, ANOVA showed a significant group effect (P = 0.0002), which indicates that the body weights differed between the groups during the intervention.
Fig. 2Body weights of the animals in the different treatment groups during the study (n = 7–8)
Effects of the atherogenic diet and test compounds on serum cholesterol and serum triglyceride concentrations
The atherogenic diet (group I) induced an almost 3-fold increase in the s-cholesterol levels but no effect was seen in the triglyceride levels. Stanol ester alone (group II) or together with sesamin (group IV) significantly attenuated the elevation of the cholesterol levels. Sesamin alone (group III) either normalize the elevation of cholesterol levels nor did it enhance the effect of stanol ester (group II versus group IV). Diet-induced changes in the triglyceride levels did not differ between groups (P = 0.89). Results are presented in Fig. 3.
Fig. 3S-cholesterol and -triglyceride concentrations before (Basal) and after (Intervention) the 4-week intervention with experimental diets and the difference between basal and Intervention levels (Difference). **P < 0.001 in comparison with Group 1 (Tukey’s Multiple Comparison Test)
Effects of the atherogenic diet and test compounds on plasma lignan profile after intervention
Presence of sesamin epimer in the diets resulted in the appearance of enterolignans in plasma, although concentrations varied greatly between animals (Table 1). Significantly increased levels of enterodiol (P = 0.00), but not enterolactone were found.
Table 1Enterolignan (enterolactone and enterodiol) values after 4-week intervention period nmol/l (Mean ± SEM, n = 8)GroupnENLaENDENL + ENDI, Control84.61 ± 0.406.11 ± 2.109.18 ± 2.14II, Stanol ester83.01 ± 0.3510.0 ± 4.6913.0 ± 4.85III, Sesamin88.01 ± 0.98569 ± 54.2**577 ± 55.0**IV, Stanol ester + sesamin76.54 ± 1.99388 ± 90.7**394 ± 91.8****P < 0.001 in comparison with Group 1 (Tukey’s Multiple Comparison Test)aENL, Enterolactone; END, enterodiol
Discussion
As one of the major components of sesame seeds, the lignan sesamin has received a great deal of interest regarding its potential as a hypocholesterolemic agent, especially after the positive results reported by Hirata et al. [7] in humans. Thereafter, different approaches have been used to confirm this observation in which different sources of dietary sesamin have been used. Studies using sesame seed-based diets provided results that are difficult to discuss since the hypocholesterolemic effect could not be primarily attributed to sesamin. As for the studies using pure sesamin supplementation, the preparations used so far contained a mixture of sesamin epimers (sesamin and episesamin in a 1:1 ratio) that they are known now to possess a different behavior both in their metabolic fate [12] and biological action [4] supporting the need to differentiate between different sesamin epimers in future studies.
A clear hypocholesterolemic effect elicited by sesamin (alone or in combination with vitamin E) was reported in studies conducted in rats [9, 13–15] or in cultured rat cells [16] and similarly, other biological effects proposed for sesamin [17–26], have also been studied using rats as experimental animal. Very few studies though, have used other animals than rats to test the effects of sesamin [27, 28]. In the recent study of Kushiro et al. [3] where sesamin was tested in different experimental animals, results for rats were confirmed, but no effect was observed in hamsters or mice. Only one study has investigated the hypocholesterolemic effect of sesamin in humans [7] finding a significant effect of dietary sesamin in the reduction of total cholesterol and LDL-cholesterol. In this study however, dietary sesamin was administered together with vitamin E and therefore the observed effect might be due to the reported synergism between this two dietary components [14] and since no separated sesamin-group was included in the study, the hypothetical action of sesamin could not be confirmed.
In order to minimize the commented variables, in the present experiment only pure sesamin epimer was used and additionally, transgenic mice were selected for they offer the most convenient model to investigate the two proposed mechanisms for the hypocholesterolemic effect of sesamin i.e. reduction of endogenous cholesterol synthesis and reduction of dietary absorption. Due to the lack of functional LDL-receptors, the animals used in this trial have a spontaneously elevated serum cholesterol concentration, and in addition, they are prone to diet-induced modulation of the serum cholesterol concentration [5, 6]. Hepatic HMG-CoA reductase is the rate-limiting enzyme in the cholesterol biosynthetic pathway and its inhibitors are very effective in lowering plasma cholesterol. Sesamin has been reported to inhibit HMG-CoA reductase in rats [13] and through this mechanism it has been proposed to reduce plasma cholesterol. However, in this study no reduction of serum cholesterol or triglyceride levels was observed in the group supplemented with sesamin alone (group III) and therefore it is concluded that sesamin does not affect HMG-CoA reductase in this animal model. Similarly, cholesterol supplementation (0.25%, w/w) for 4 weeks increased plasma total cholesterol levels approx. 3-fold in the absence of a functional LDL receptor. However, the introduction of stanol ester in the atherogenic diet significantly reduced the total plasma cholesterol level, independently of the presence of sesamin. It is known that plant stanols inhibit the absorption of dietary cholesterol and the reabsorption (enterohepatic circulation) of endogenous cholesterol from the gastrointestinal tract [29]. It seems though that sesamin does not affect dietary cholesterol absorption in this animal model.
The metabolism of sesamin seems to play an essential role in its further biological action. It has been recently suggested that in humans, sesamin is absorbed through the portal vein reaching the liver where extensive metabolism takes places generating demethylenated (catechol) derivatives secreted as glucuronidates via the bile [26]. The same metabolites were found after in vitro fermentation of pure sesamin standard with human fecal inoculum and furthermore extensive in vivo conversion to enterolactone was reported after sesame seed supplementation in humans [11]. We hypothesized then that the absorption of sesamin could certainly take place via the portal vein and that transformation to catechol derivatives could be possible, as well as further absorption to the general circulation where they may exert the biological actions proposed for sesamin. Nevertheless, in this study as in our previous trial [11], extensive conversion of sesamin to enterolignans was observed suggesting that non-absorbed sesamin and possibly biliary catecholic sesamin derivatives undergo metabolism by gut microflora leading to the production and further absorption of enterolignans. Postprandial levels of sesamin in plasma differed considerably between rats [12] and humans [11], suggesting a different metabolic pathway depending on species.
So far, our data suggest that a limited absorption or a too rapid enterohepatic circulation of sesamin could be the explanation for the lack of antihypercholesterolemic effect in mice/hamsters and possibly in humans but this hypothesis needs to be confirmed. Furthermore, the characteristics of our animal model did not allow to study a third mechanism for cholesterol reduction that sesamin could use; the increment of LDL-receptor activity. This hypothetical mechanism has been suggested for other dietary components [30], and it is a likely explanation of the negative results reported in this paper.
In conclusion, possible confounding factors have been minimized with the use of a specific animal model and the selection a pure sesamin epimer. It can be therefore stated that sesamin does not seem to affect cholesterol biosynthesis or absorption in mice. The so far contradictory results hinder the extrapolation to humans, and only a clinical trial with separate epimeric forms and the elucidation of the complete metabolic pathway of sesamin in humans will contribute to clarify the possible utilization of sesamin as a hyocholesterolemic agent.

sesamin
cholesterol
triglycerides
lignans
stanol ester
enterolignans