BioMed Research International

BioMed Research International / 2014 / Article
Special Issue

How Microgravity Affects the Biology of Living Systems

View this Special Issue

Review Article | Open Access

Volume 2014 |Article ID 928507 |

Claudia Ulbrich, Markus Wehland, Jessica Pietsch, Ganna Aleshcheva, Petra Wise, Jack van Loon, Nils Magnusson, Manfred Infanger, Jirka Grosse, Christoph Eilles, Alamelu Sundaresan, Daniela Grimm, "The Impact of Simulated and Real Microgravity on Bone Cells and Mesenchymal Stem Cells", BioMed Research International, vol. 2014, Article ID 928507, 15 pages, 2014.

The Impact of Simulated and Real Microgravity on Bone Cells and Mesenchymal Stem Cells

Academic Editor: Mariano Bizzarri
Received04 Apr 2014
Revised06 Jun 2014
Accepted06 Jun 2014
Published10 Jul 2014


How microgravity affects the biology of human cells and the formation of 3D cell cultures in real and simulated microgravity (r- and s-) is currently a hot topic in biomedicine. In r- and s-, various cell types were found to form 3D structures. This review will focus on the current knowledge of tissue engineering in space and on Earth using systems such as the random positioning machine (RPM), the 2D-clinostat, or the NASA-developed rotating wall vessel bioreactor (RWV) to create tissue from bone, tumor, and mesenchymal stem cells. To understand the development of 3D structures, in vitro experiments using s- devices can provide valuable information about modulations in signal-transduction, cell adhesion, or extracellular matrix induced by altered gravity conditions. These systems also facilitate the analysis of the impact of growth factors, hormones, or drugs on these tissue-like constructs. Progress has been made in bone tissue engineering using the RWV, and multicellular tumor spheroids (MCTS), formed in both r- and s-, have been reported and were analyzed in depth. Currently, these MCTS are available for drug testing and proteomic investigations. This review provides an overview of the influence of on the aforementioned cells and an outlook for future perspectives in tissue engineering.

1. Introduction

It is well known, that microgravity influences different biological systems like bone and muscle as well as the heart and brain, and it enhances cancer risk [1]. During their stay at the MIR, astronauts and cosmonauts did show a distinct loss of bone mineral density in the lumbar spine, the pelvis, and the proximal femur [2], and the extent of bone loss varied up to 20% [3].

As it is not feasible to gather enough material from astronauts to do in-depth investigations, another device has been developed for the International Space Station (ISS), the mice drawer system (MDS), as a facility to study long-time influence of radiation on the biology and behavior of mice. Tavella et al., for example, report an altered bone turnover in different strains of mice which were kept on the ISS for 91 days. This resulted in bone loss due to increased bone resorption and a decreased bone deposition [4].

While the past biological, physiological, and medical research nearly exclusively focused on investigating the biochemical processes of living cells and organisms, more and more attention was paid to the biomechanical properties and mechanical environment of cells and tissues during the last decades. When culturing cells on Earth, they usually settle on the bottom of the culture flask, forming two-dimensional (2D) monolayers. A three-dimensional (3D) growth, more resembling the tissue environment found in living organisms, is prevented by the presence of the gravitational field. For a scaffold-free 3D tissue growth, it is therefore necessary to circumvent this problem by effectively eliminating the influence of the gravitational pull during cultivation. One of the byproducts of various space flight endeavors is the possibility to perform long-term near-weightlessness or microgravity () experiments [5, 6]. In a environment, cells will not settle like on Earth. This provides an increased opportunity for freely floating cells to interact with each other and develop 3D structures [7].

2. Space Flights for Cell-Biological Experiments

Long-term orbital space flight experiments are, however, not trivial. Flight opportunities are very scarce and the costs of hardware development are high. Furthermore, science is not always a priority in space flight activities. Such preconditions are delaying the advancement of research in areas such as cell biology and tissue engineering disciplines, which could profit tremendously from more frequent research options in a real microgravity (r-) environment.

Some researchers recently pointed out that osteoblasts undergo a disintegration of their cytoskeleton, which may explain dramatic changes in size and shape of the cells and their surface specializations [47]. Also, other studies have been performed using the ISS or space shuttle flights to learn more about the behavior of bone cells in space [48], but flight opportunities are sparse, and, therefore, other platforms had to be elucidated.

It is due to the aforementioned limitations that, over the years, various devices have been developed in an attempt to reduce the impact of gravity and simulate a near-weightlessness environment (s-) on Earth. From a physical point of view, gravity is a force exhibiting both magnitude and direction. Therefore, the influence of gravity can be reduced by either manipulating magnitude or direction. An orbital space flight as on the ISS is physically identical to a free-fall. Here, the gravitation acts in a perpendicular manner on the spacecraft’s velocity vector, effectively changing its direction constantly but not affecting its magnitude. Free-fall is also found when using sounding rockets, which provide r- during a time span of up to 15 minutes. On Earth, r- can also be attained, although only for periods in the range of seconds, in drop towers, and during parabolic flights missions [49, 50]. Although time periods of seconds or minutes limit their use for tissue engineering studies, such periods can be useful to explore various intra- and intercellular processes, responsible for gene expression and protein content changes which can be observed after only a few hours of culturing cells in [4951].

3. Devices Simulating Microgravity on Earth

In this respect, we should mention an instrument that was introduced by the European Space Agency (ESA) in the early nineties, called the free fall machine (FFM) [52]. This instrument was specifically developed for biological experiments and could generate a free fall for a period of about 800 ms with an intermediate “bounce” of ~20 g for around 50 ms. The paradigm of the FFM is that cells might not be sensitive to the relatively short period of 50 ms of hypergravity, while they experience the relatively longer period of free-fall. Long-term experiments (hours, days), which might be useful for tissue engineering studies, could be performed on this platform. However, thus far, only two studies were published using the FFM, one investigating Chlamydomonas [53] and another one researching T-lymphocytes [54]. The Chlamydomonas study showed similar results to what was found in real space flight while the T-lymphocytes experiments did not. Considering the very limited number of studies performed on this ground-based device, the FFM still might deserve some more exploration.

Levitating magnets are also used to produce s- on Earth. Such systems compensate the magnitude of the gravity vector by preventing sedimentation of relatively heavy structures, like cells, by the application of a high gradient magnetic field. This principle was first described for biological systems by Berry and Geim in 1997 [55], who demonstrated that a toad could be levitated and survive while exposed to a 16 Tesla magnetic field. Various experiments in cell biology have made use of such systems [5658]. The magnetic field acts on individual molecules and atoms within a cell, based on their magnetic susceptibility, preventing them from sedimentation. However, the magnetic field as such confounds possible s- effects. The direction of the field might force (bio-)polymers into a certain orientation. Different polymers within a cell or on the cell membrane have different susceptibilities, possibly producing artifacts by forcing polymers into specific arrangements, which may not reflect the actual physiological situation [5961]. Superconducting high gradient magnets are especially capable of performing long-term experiments and might be useful in the area of tissue engineering [6264]. In this context, another promising technique should be mentioned. This method is the use of magnetic particles for 3D cell cultures. It is not based on a high-gradient magnetic field, but on ferromagnetic particles attached to cells, which can subsequently be levitated by a conventional magnet facilitating the formation of 3D structures [65, 66].

Another option is to manipulate the direction of the gravity vector with respect to the sample. The reduction of the gravitational impact on biological systems by constantly changing its orientation was shown first in experiments by the German botanist von Sachs in 1879, growing Lepidium sativum and Linum usit [67]. He constructed a slowly rotating system and named it a clinostat, in which, for example, a plant can be placed horizontally and rotated around its longitudinal axis. In doing so, the gravity vector stimulus is constantly changing its impact angle on the sample. As a result, a plant grows straight without the characteristic gravitropic curvature seen when the plant is placed horizontally and not rotating. Based on these initial studies, other rotating systems like the fast rotating clinostat have been developed.

The initial clinostats were rotating relatively slowly in a range from one rotation per couple of hours up to a maximum of about 10 rpm. This is adequate for relatively “solid samples” such as plants, but too slow for cell culture systems that involve a large liquid phase. In a biphasic system, that is, a liquid with particles (cells) both of different density, the heavy particles tend to settle. Rotating such a system around a horizontal axis keeps the heavy particles in suspension. This phenomenon depends mainly on the relative density of the liquid and the particles, the viscosity of the liquid, the rotation speed, and the diameter of the rotated container. When a cell is in a static vessel and the vessel is rotated by 90°, the cell will settle in the direction of the gravity vector. One can repeat this for a full 360° and upon an increase in the frequency of rotation, the traveling distance of the cell decreases. If this rotation is performed constantly with increased speed, we finally end up rotating a cell around its own axis. Such a controlled rotation not only applies to the cells, but also its surrounding boundary liquid phase [68].

Another well-known device to simulate is the so-called random positioning machine (RPM), a 3D clinostat [69] consisting of two frames, each driven by a dedicated motor. This allows a randomized movement of both frames, independent of each other [6974]. One of the advantages of the RPM is its size, as cell culture flasks can easily be mounted on it, so it is possible to work with quite large liquid volumes. This ranges from regular T25 flasks [75, 76] to multi-well plates [77], flasks on slides [78], or more dedicated devices [79]. As cells move freely within the liquid, they usually interact with each other and form multicellular spheroids.

The best simulation of is achieved in the rotation center of the two axes, which limits the preferred volume size of the samples. Depending on the speed of rotation and the distance from the center, an acceptable residual gravity can be obtained in the order of 10−4 g by a maximum angular velocity of 60° s−1 at a radial distance of 10 cm [70]. Earlier RPM models had no possibility to add constituents during the experiment, but newer models have been developed to enable fluid management during rotation [73, 74]. RPMs are commercially available by Mitsubishi Heavy Industries (Kobe, Japan) and Dutch Space (Leiden, The Netherlands), while various academic groups developed similar systems on their own [8084] (Figure 1).

The rotating wall vessel (RWV) prevents cells from settling via a constant rotation. It has been developed by NASA [85] and is now commercially available through Synthecon Inc. (Houston, TX, USA). Basically, RWVs consist of a slow rotating, relatively large liquid filled container (vessel). The rotation speed has to be adapted to the specific weight of the cells, the fluid density, and viscosity. The cells and tissues in the RWV are constantly falling within the fluid. The settling velocity and direction combined with the rotation of the fluid create spiral trajectories within the vessel [86]. This motion of the sample relative to the fluid generates fluid shear forces on a particle surface ranging from 180 to 320 mPa (1.8–3.2 dyne/cm2) for 50 μm beads [87], ~500 mPa (5 dyne/cm2) with 3D aggregates of BHK-21 cells [88] to 520–780 mPa (5.2–7.8 dynes/cm2) for a 200 or 300 μm spherical object [89]. Over the years, various models based on the initial RWV have been developed, differing in vessel geometry, aspect ratio, and gas supply, such as the slow turning lateral vessel (STLV) [90], the high aspect ratio vessel (HARV) [91], or the rotating-wall perfused vessel (RWPV) [92].

Hence, it can be concluded that annulling the gravity forces, which pull the cells constantly towards the Earth, deliver the ultimate trigger to eukaryotic cells to leave a cell monolayer and assemble in 3D aggregates [5].

It is still unknown which cellular and biochemical mechanisms are involved in the altered signal transduction and in the change of the cellular growth behavior.

4. Transition from Two- to Three-Dimensional Cell Growth

A few publications appeared in the literature in recent years, providing some clues for understanding the weightlessness-induced transition from two- (2D) to three-dimensional (3D) cell growth.

Several signaling pathways are affected by annulling gravity forces in the cell interior [93]. However, it is unknown which of these signaling pathways contribute to the formation of three-dimensional aggregates. When endothelial cells form tubes, the nitric oxide signaling pathway appears to be affected [94]. Siamwala et al. reported that iNOS (inducible nitric oxide synthase) acts as a molecular switch, which controls whether the effects of on vascular endothelial cells induce angiogenesis via the cyclic guanosine monophosphate (cGMP)-PKG-dependent pathway [94]. iNOS is upregulated in HUVEC by a mechanism dependent on suppression of AP-1, after clinorotation of the cells [95]. In addition, the endothelial nitric oxide synthase is phosphorylated by phosphoinositide 3-kinase under weightlessness, simultaneously with Akt [96]. The organoid formation by PC12 pheochromocytoma cells in a RWV bioreactor is accompanied by prolonged activation of the ERK, p38, and jnk signaling pathways [97].

3D cell culture techniques have attracted much attention, not only among biologists, but also clinicians interested in tissue engineering [98, 99] of artificial vessels [100104] or cartilage [105108]. Moreover, osteoarthritis and cartilage trauma occur in patients with a high incidence, but current treatment methods are still limited [109]. Even a minor injury to articular cartilage may lead to progressive damage and degeneration [110].

5. Tissue Engineering of Bone

Bone loss has been documented for many years in (1-2% a month). Increased bone loss and risk of fractures is an identified risk in the bioastronautics critical roadmap for long-term cosmic missions to the moon and mars. In vitro drug screening both in 1 g, and in artificial gravity is essential to adequately address countermeasures for bone loss. Bone loss in is the second most important risk to space missions [5, 6].

Exposure to the environment of space causes astronauts to lose calcium from bones [5, 6]. This loss occurs because the absence of Earth’s gravity disrupts the process of bone maintenance in its major function of supporting body weight. Exposure to the environment of space causes men and women of all ages to lose up to 1% of their bone mass per month due to disuse atrophy, a condition similar to osteoporosis. It is not yet clear whether loss in bone mass will continue as long as a person remains in the environment or level off in time.

There are, indeed, four major bone cell types, and each of them seems to be influenced by . Bone mesenchymal stem cells (MSC) are able to differentiate into adipocytes, osteoblasts, and osteoclasts. Proliferation and differentiation are very sensitive to , as the lack of gravity in space can reduce mechanical stress, leading to a decreased rate of osteogenesis and an increased adipogenesis rate [111]. As the signaling pathways involved in MSC differentiation form a complicated network, it has been found that the reduction in the osteogenesis of MSCs in the presence of is mediated by a decrease in the integrin/mitogen-activated protein kinase (MAPK) signaling pathway [112], as well as RhoA and cytoskeletal disruption [113].

Osteoblasts are derived from MSCs, but in the differentiation does not function properly, and the resulting bone loss has been attributed to osteoblasts due to their (1) reduced proliferation and activity, (2) reduced differentiation, and (3) decreased responsiveness to bone-related factors in the microenvironment [114]. Observations have also been made regarding the cytoskeleton of osteoblasts; there is growing evidence that the cytoskeleton is closely connected to nuclear morphology and function [115]. The enlarged nuclei observed in flight osteoblasts could be a result of cytoskeletal disruption [116].

Osteocytes regulate bone resorption and formation and are considered the terminal differentiation stage of osteoblasts. The osteocytes in cortical bone and periosteum degenerated after a 12.5-day flight in space on the Cosmos Biosatellite [117]. Osteocyte apoptosis has been observed after a 2-week flight, increasing the number of functionally active osteoclasts [118]. Apoptotic osteocytes are essential for the initiation of bone remodeling, but it is the neighboring nonapoptotic osteocytes that produce proosteoclastogenic signaling [119]. Osteocytes seem to be the key effectors of induced bone loss [120].

Osteoclasts are bone-resorbing cells, and their differentiation seems to be enhanced in [121]. This could be another explanation of bone-loss in space.

The mystery, for the moment, is what signals permit bone tissue to adapt to a weightless or an Earth (1 g) environment. Researchers do not yet know whether the biomechanical stimuli that are changed by directly affect osteoblast and osteoclast function or if other physiological factors such as hormone levels or poor nutrition contribute to bone loss. NASA investigators are studying gravity-sensing systems in individual bone cells by flying cultures of these cells on the space shuttle and observing how they function. Discoveries made in the course of space biomedical research on bone are already contributing to a better understanding of osteoporosis and the treatment of bone mass loss on Earth as well as in space. The single most important contribution that NASA research has made to the understanding of bone deterioration in osteoporosis is heightened awareness of the importance of gravity, activity, and biomechanics—that is, the mechanical basis of biological activity— in bone remodeling.

Mechanical forces—the action of energy on matter—appear to coordinate bone shaping processes. The standard theory of bone remodeling states the body translates mechanical force into biochemical signals that drive the basic processes of bone formation and resorption. Aging, especially in postmenopausal women, and exposure to uncouple bone resorption and formation. When this uncoupling occurs, formation lags behind resorption, and the result is bone loss.

Researchers are not yet certain whether bone resorption speeds up or the bone formation slows down, though recent experimentation in space indicates that might somehow affect both processes. Progress in developing methods of preventing or treating disuse atrophy and osteoporosis depends on better understanding of the mechanisms that cause the problem. Determining how the body translates mechanical loading (physical stress or force) into the signals that control bone structure may reveal how aging, inactivity, and space flight uncouple bone formation and resorption. Only in the absence of gravity can we determine the influence of weight and stress on bone dynamics.

By studying what mechanisms translate mechanical stress on bones into biochemical signals that stimulate bone formation and resorption, space life scientists may be able to determine how to maintain bone mass. Researchers do not yet know exactly what type and amount of exercise, hormones, or drugs might prevent bone loss or promote bone formation. However, some combination of sex hormones, growth hormones, and exercise seems to be the key to preventing bone mass loss associated with chronological aging and postmenopausal hormone changes on Earth.

Bone is made up of several different cell populations. Osteoclasts are responsible for the breakdown of mineralized bone, in preparation for bone remodeling. In contrast, the osteoblasts synthesize mineralized bone in the remodeling process. The goal of this project is to develop an “in vitro” three-dimensional, cellular model of osteoclasts and osteoblasts (human and rodent) cultured together in analog culture conditions to identify the underlying biomarkers related to bone loss in and the cellular mechanisms involved in bone resorption. The NASA rotating-wall vessel (RWV) permits the growth of mixed cell cultures for much longer periods than traditional culture methods. This would set the stage for development of countermeasure strategies for bone loss in space as well as in osteoporosis and rheumatoid arthritis which are increased health risks on Earth. Professor Sundaresan and collaborators [122124] have developed a 3D cell culture bone tissue model using a specialized rotating-wall vessel culture system to address a more physiologically relevant model to the human body. The use of the cells by themselves also eliminates confounding variables such as neuroendocrine stress found in vivo (Figure 2(a)).

The human body needs a framework to withstand gravity. This framework is given by the skeletal system. During long-term space missions, bone loss has been reported in astronauts at a rate that is both substantial and progressive with time spent in [125128]. But what is the reason for this massive bone loss? Some studies suggested that this effect might be attributed to increased resorption in load-bearing regions of the skeleton [129131], and evidence of a decrease in bone formation had also been described. For example, the loss of bone in is about 10 times greater than the bone mineral density loss per month of postmenopausal women on Earth, who are not on estrogen therapy [132135]. The loss of bone mineral density in a six-month mission appeared to be reversible in 1000 days after return to Earth [136, 137], but changes in the bone structure are irreversible and seem to mimic changes in the elderly [137].

Until now there are still knowledge gaps on the mechanism of bone loss, especially on the molecular and cellular mechanisms, also the question of fracture repair arises. Moreover, more information is needed on the influence of radiation, hormones, and fluid shifts.

Investigations in humans and animals are quite difficult due to the lack of long-term flight opportunities, the absence of animal housing facilities in space, and the problem of material collection from returning astronauts. Thus, other possibilities have to be sought in order to investigate bone. So far, most commonly used are bone cell culture experiments, which are a viable opportunity for investigating cells in 3D, acting as tissue like samples while they are cultivated under conditions of weightlessness. However, 3D embryonic bone tissue cultures have been used in the past and show a clear decrease in matrix mineralization, in mineralizing cartilage and by osteoblasts, combined with an increased mineral resorption by osteoclasts [138].

Besides this, tissue engineering is a very up-to-date topic. The ultimate goal is to generate functional 3D constructs, which can be used as replacement organs or structures with normal function or serve for in vitro studies [5, 139]. Bone replacement, especially, is quite difficult, as large bone defects usually require reconstructive surgery to restore function [140]. Up to date, the treatment includes autograft or allograft transplantation and the use of synthetic materials [141]. While autograft transplantation is the preferred treatment, it suffers from limited supply and donor site morbidity [142]. As the autogenous origin of cells prevents potential immune rejection, the amount of bone marrow suitable for transplantation is limited. New techniques have been developed, allowing selection of bone marrow osteoprogenitor cells and expanding them in culture, so that a large amount of transplantable cells can be generated after only one biopsy [143145].

In principle, culturing bone cells is not that easy. A combination of osteoconductive matrices, bone-forming cells, and osteogenic growth factors is needed for the engineering of bone tissue [146]. The first important factor is the cell type. Osteoblasts are in a close to mature stage, showing a low proliferative potential. Mesenchymal stromal cells represent a proliferating and undifferentiated cell source, but their availability is limited [147, 148]. An option to increase their lifespan in vitro is the overexpression of human telomerase reverse transcriptase (hTERT). The second factor is an ideal scaffold, which possesses mechanical properties comparable to bone. It should support cell adhesion and should be biodegradable to facilitate natural bone remodeling [146]. As of now, different studies have shown the advantages and disadvantages of several types of scaffolds like chitin, gelatin, poly(lactic acid), poly(glycolic acid), poly(lactic acid-co-glycolic acid), polycaprolactone, hydroxyapatite, coral, and so forth. Several in vitro studies revealed an ideal scaffold pore size for osteoblasts from 200 to 400 μm [149, 150]. It is important to recognize that the scaffold architecture influences the distribution of shear stress, the range of mechanical stimuli, as well as the proliferation and differentiation of osteoprogenitor cells [151, 152].

To simulate an ideal in vivo situation for in vitro cells, specific cytokines and growth factors are necessary. For bone morphogenesis, the bone morphogenetic proteins (BMP), which belong to the transforming growth factor beta (TGF-) superfamily, are essential [153]. Currently, only BMP-2 and -7 are commercially available, so alternatives to stimulate osteoprogenitor cells by growth factors are required. It has been reported that autologous platelet-rich plasma is an effective bioactive supplement, as it contains osteogenic and angiogenic growth factors [154].

Several different bioreactor systems are already available for bone tissue engineering. A well-known and simple system is the spinner flask bioreactor. Convective forces are provided by a stirrer and the medium flows around the cells. The emerging shear stress is not applied homogenously, as there appears to form a gradient in the flask [146]. This factor certainly needs to be considered when conducting studies with the spinner flask system.

Other suitable instruments are rotating bioreactor systems, for example, the RWV. It has been used with different kind of bone cells, which are often grown with the help of microcarriers [8, 155] or scaffolds [811, 15, 155]. The high aspect ratio vessel (HARV) [91] was used by Lv et al. [12] to engineer tissue on poly(lactic acid glycolic acid)/nano-hydroxyapatite composite microsphere-based scaffolds.

Some researchers used bone marrow mesenchymal stem cells for their investigations. Jin et al. [16] were able to transplant RWV-grown bone constructs in cranial bone defects of Sprague-Dawley rats and found them to be more effective in repairing the defects than the 1 g controls after 24 weeks. Moreover, a 3D environment as in a rotary cell culture system enhanced osteoblast cell aggregation and mineralization [13]. Preosteoblasts cultured in a RWV could be engineered into osseous-like tissue [14].

6. Mesenchymal Stem Cells and Microgravity

Mesenchymal stem cells (MSCs) are cells capable of long-term proliferation and differentiation into various stromal tissue cell types. The state of MSCs rests on the cellular microenvironment and several soluble factors. In addition, gravity can influence MSC features. Disuse, as encountered during long-term bed-rest or space travel, and the accompanying absence of mechanical stimuli lead to an inhibition of osteogenesis and simultaneously to an induction of adipogenesis in MSCs. Hence, it is crucial to provide a proper mechanical stimulation for cellular viability and osteogenesis, particularly under unusual conditions.

In 2004, Merzlikina et al. [27] studied the effects of prolonged clinorotation on cultured human MSC morphology, proliferation rate, and expression of specific cellular markers. After exposure of the cells to clinorotation for time frames from 1 h to 10 days, it was shown that the proliferative rate decreased in the experimental cultures as compared to cells growing under normal conditions. Clinorotated MSCs seemed more flattened and reached confluence at a lower cell density, which advocates that cultured hMSCs sense the changes in the gravity vector and respond to s- by altered functional activity. The group around Myoui [28] examined whether gravity-induced stress is linked to osteoblast differentiation and function. Rat marrow mesenchymal cells (MMCs) were cultured in pores of interconnected porous calcium hydroxyapatite (IP-CHA) for 2 weeks on a 3D clinostat. In MMCs subjected to s-, the marker of osteoblastic differentiation alkaline phosphatase activity was decreased by 40%, compared to the control group. Also, the clinostat group exhibited less extensive extracellular matrix formation than the control group. The implantation of the IP-CHA/MMC composites in syngeneic rats showed that bone formation was significantly lower for the clinostat group than for the control group. Yuge et al. [29] also used a 3D clinostat for their experiments on the proliferation behavior of hMSCs. The proliferation rate of the cells of the clinostat group was elevated almost 3-fold in comparison to the control group, and the number of hMSCs double-positive for CD44/CD29 or CD90/CD29 in the clinostat group after 7 days in culture increased 6-fold. The hMSCs cultured in a 3D-clinostat were still able to differentiate into hyaline cartilage after transplantation into cartilage defective mice and displayed the strong proliferative characteristic of stem cells, thus, showing that s- may be used to expand stem cell populations in vitro. In contrast to these findings, Dai et al. [24] reported in 2007 that simulated by a clinostat inhibited population growth of bone marrow mesenchymal stem cells (rBMSCs) and their differentiation towards osteoblasts. The cells grown on the clinostat were arrested in the G(0)/G(1) phase of cell cycle, and growth factors, such as insulin-like growth factor-I, epidermal growth factor, and basic fibroblast growth factor had only a slight stimulatory effect compared to the static control group. Gershovich and Buravkova’s [17] work supports this hypothesis. After 20 days of clinostat-exposure, the proliferative activity of hBMCs was reduced, whereas it increased the number of large flat cells in the culture and stimulated migration activity of cells. In 2009, Gershovich and Buravkova [30] demonstrated the effects of s- by clinostat and RPM on the interleukin production by hBMSCs and MSC osteogenous derivatives. 20-day exposure on a clinostat increased the interleukin-8 (IL-8) content 1.4 to 3.2 times in the culture medium, while the average increase of IL-production on the RPM amounted to 1.5–6 times (10 days) and 1.6–2.1 times (20 days), respectively. This suggests that results of s- vary by the use of different modeling systems. rMSCs grown in a clinostat demonstrate that s- can boost the differentiation of MSCs into neurons, as demonstrated by Chen et al. [156] In s-, neuronal cells derived from rMSCs were found to express higher microtubule-associated protein-2 (MAP-2), tyrosine hydroxylase (TH) and choline acetyltransferase (CHAT). Furthermore, the excretion of neurotrophins such as nerve growth factor (NGF), brain derived neurotrophic factor (BDNF), or ciliary neurotrophic factor (CNTF) was increased. In comparison to 1 g controls, neuronal cells from the s- group generated more mature action potentials and displayed repetitive action potentials. This might benefit the search for new strategies for the treatment of central nervous system diseases.

Zayzafoon et al. [18] demonstrated that s- inhibits the osteoblastic differentiation of hMSC and induces the development of an adipocytic phenotype. In the effort of understanding space flight-induced bone loss, the group used the rotary cell culture system (RCCS) to model and determine its effects on osteoblastogenesis. Human MSCs were cultured and osteogenic differentiation was induced before the initiation of s-. As a result, the important mediator of adipocyte differentiation, peroxisome proliferator-activated receptor gamma (PPARgamma2), and adipsin, leptin, and glucose transporter-4 was highly expressed. These changes were not adjusted after 35 days of readaptation to normal gravity. Moreover, decreased ERK- and increased p38-phosphorylation pathways, known to regulate the activity of runt-related transcription factor 2 and PPARgamma2. These results were supported by Saxena et al. [19] in 2007, who demonstrated that s- inhibited osteoblastogenesis and increased adipocyte differentiation in hMSCs incubated under osteogenic conditions using the RCCS. They could show that a reduced RhoA activity and cofilin phosphorylation, disruption of F-actin stress fibers, and decreased integrin signaling through focal adhesion kinase were involved in this process. Meyers et al. [20] also investigated the effects of s- on integrin expression and function in hMSCs, since a reduced osteoblastic differentiation might be caused by impaired type I collagen (Col I)-integrin interactions or a reduction of integrin signaling. Culturing of hMSCs for 7 days in s-, lead to reduced expression of Col I, while Col I-specific alpha2 and beta1 integrin protein expression increased. However, autophosphorylation of adhesion-dependent kinases, focal adhesion kinase (FAK) and proline-rich tyrosine kinase 2 (PYK2) was significantly reduced. These findings indicate that a reduction in osteoblastogenesis in s- is, at least in part, caused by a reduced integrin/MAPK signaling. The group around Duan [16] studied the relationships between the composition and mechanical properties of engineered bone constructs. BMSCs were grown for 15 days on ceramic bovine bone scaffolds in different environments, namely, static flasks and the RWV. DNA content and alkaline phosphatase (ALP) were higher for cells grown on the RWV. After transplantation into Sprague-Dawley rats with cranial bone defects, the bone constructs engineered on the RWV repaired the defects better and showed histologically better bone connection.

Sheyn et al. [21] evaluated the effect of s- on all genes expressed in hMSCs, with the hypothesis that many important pathways are affected during culture on a rotating wall vessel system. The analysis of gene expression by use of whole genome microarray and clustering showed that 882 genes were downregulated and 505 genes were upregulated after exposure to s-v. A multitude of genes belonging to cell compartment, biological process, and signaling pathway clusters were modulated, as identified by gene ontology clustering. Significant reductions in osteogenic and chondrogenic gene expression and an increase in adipogenic gene expression were shown and could be validated by a parallel adipogenic differentiation assay. In order to investigate the effects of s- on chondrogenic differentiation of human adipose-derived MSCs (ADSCs), Yu et al. [22] cultured cells on a RCCS in pellets with or without the chondrogenic growth factor TGF-. Analysis of real-time PCR and histological results demonstrated that s- has a synergistic effect on chondrogenesis with TGF-. The p38 MAPK pathway was activated by TGF- alone and was additionally stimulated by s-. Inhibition of p38 activity with SB203580 suppressed chondrocyte-specific gene expression and matrix production. This indicates that the p38 MAPK signal mediates s--induced chondrogenesis of ADSCs. In MSCs cultured during chondrogenic induction in a rotating culture, combined with polyglycolic acid (PGA), mRNA and proteins of collagen type II and aggrecan were significantly more expressed in the s- rotating culture group than the static culture group, as reported by Wu et al. [25]. Zhang et al. [26] described that MSCs spread out in a spindle shape when cultured in normal gravity, while they become unspread and round under s-. Also, under s-, their cytoskeleton fibers are being reorganized. The function of MSCs was affected by these morphological changes, transmitted through the activity of RhoA. To test the hypothesis that s- has the capacity to offer a novel choice in the stimulation of neovascularization, MSCs were cultured under s- stimulation followed by VEGF differentiation. The responses revealed that MSCs were differentiated into endothelial-like cells after 72 h incubation and were able to form a capillary network. Their endothelial differentiation potential improved compared with the static control group.

Another approach of modeling in hMSCs is the use of a large gradient high magnetic field (LGHMF) produced by a superconducting magnet. Shi et al. [64] analyzed the effects of LGHMF- on survival, cytoskeleton and osteogenic potential of hMSCs. Results showed that the LGHMF- treatment disrupted the cytoskeleton of hMSCs, a LGHMF- treatment for 24 h led to cell death. LGHMF- treatments in early stages of osteogenic induction resulted in suppression of osteogenesis of hMSCs. The suppression intensity was reduced gradually as the treatment stage of LGHMF-MG was postponed. A LGHMF- treatment during the ending-stage of osteogenic induction had no visual effect on osteogenesis of hMSCs, which indicates that LGHMF- affects the initiation of osteogenesis.

Furthermore, a study of Uddin and Qin [23] examined the effects of low intensity pulsed ultrasound (LIPUS) on the osteogenic differentiation of adipose-derived human stem cells (Ad-hMSC) under s- conditions. Microgravity was simulated in a 1D clinostat and treated with LIPUS at 30 mW cm−2 for 20 min day−1. Hypothetically, the application of LIPUS to s- cultures would restore osteogenesis in Ad-hMSCs. The results showed significant increases in ALP, OSX, RANKL, and RUNX2 and decreases in OPG gene expression in LIPUS treated SMG cultures of Ad-MSC compared to nontreated cultures. LIPUS also restored OSX, RUNX2, and RANKL gene expression in osteoblast cells. s- significantly reduced ALP positive cells by 70% () and ALP activity by 22% (), while LIPUS treatment restored ALP positive cell number and activity to equivalence with normal gravity controls. Extracellular matrix collagen and mineralization was assessed by Sirius red and Alizarin red staining, respectively. s- cultures showed little or no collagen or mineralization, but LIPUS treatment restored collagen content to 50% () and mineralization by 45% () relative to s-—only cultures.

7. Multicellular Tumor Spheroids

3D growth of tumor cells creating MCTS in vitro has been observed in various tumor cell lines including thyroid and colorectal cancer [31]. MCTS mimic the growth of solid tumors and represent a simple model, approaching some of the characteristics found in vivo including physiological characteristics such as multicellular architecture and natural barriers of mass transportation. Therefore, the use of MCTS as an in vitro tool for testing anticancer drugs has gained significant interest as MCTS potentially provide a more reliable model for drug testing compared to single layer adherent cell cultures. During the approval process of drugs before clinical testing in trials, the mechanisms of delivery and the effectiveness of the drugs must be determined. The first steps of preclinical drug testing are typically carried out using adherent cell formats growing in two dimensions [157]. However, the outcome of such investigations in two-dimensional cell systems is often very different from what is observed in a whole-body situation. This makes it difficult to draw clear conclusions of the drug properties anticipated in vivo. In terms of drug delivery, a spheroid test platform has inherent advantages, providing a natural barrier resembling the natural tumor environment. Spheroids of a particular size exhibit certain gradients of oxygen and nutrition [3537, 158, 159]. Spheroids larger than 400–500 μm in diameter show characteristics of layered structures with a hypoxic core, consisting of necrotic cells surrounded by quiescent cells and an outer layer of proliferating cells [38, 160, 161]. Hence, 3D tumor cell systems are a valuable tool for studying drug delivery and the response and metabolism of hypoxic tumor cells to cancer therapy. Fang et al. reported that multicellular spheroids of primary human colon cancer cells were resistant to chemotherapy-induced apoptosis and retained the expression of colon cancer marker CD133, mimicking colorectal cancer [162]. Were these cells grown under normal conditions, they did not retain these characteristics. Size control of MCTS is a major challenge in obtaining uniform and reliable high throughput test systems; various techniques such as forced aggregation techniques, micro textured surfaces, and porous 3D scaffolds are being employed to solve these issues [163166]. There are several conventional methods for generating 3D aggregates of cancer cells, including NASA rotary cell culture systems, hanging drops, and culturing of cells using nonadherent surfaces [166168]. Conditions of s- using the RPM (Figure 2(b)) or the HARV have been shown to induce the growth of MCTS without the use of scaffolds in several types of human cancer cells [3133, 39, 169]. The molecular switches initiating s--induced spheroid formation are still unknown. Several changes in morphology and gene expression profiles have been observed in follicular thyroid cancer cells, grown under s- conditions with the main features involving changes in the ECM and early induction of apoptosis [39, 40]. Signaling between exogenous ECM and tumor surface receptors has long been thought to be an essential component in regulating the tumorigenic phenotype in 3D cultures. These phenomena have been demonstrated in studies showing that blocking specific ECM-integrin signaling can cause a shift in the malignant potential of tumor cells, leading to a more benign phenotype [40, 170]. In an attempt to identify gravity sensitive genes responsible for MCTS formation, Grosse et al. [41] identified 487 transcripts, which were differently regulated after 24 h of s- in comparison to the ground control. Comparing adherent cells with MCTS under s- conditions revealed significant differences in terms of growth patterns and signaling. Interestingly, the rate of apoptosis was increased in adherent cells compared to MCTS, indicating that the early phase induction of apoptosis may be concomitant with the transition of cells shifting from 2D to 3D growth. Several NF-κB-driven genes, involved in the regulation of tumor invasion, were upregulated by s- in adherent cells, highlighting that s- initiates distinct adaptive mechanisms in the cells.

8. Summary

The development of tools like the RPM or RWV gave new impacts in the field of tissue engineering (Table 1). Growing cells in simulated or real weightlessness, for example, on the RPM, might be a highly promising new technique to generate tissue constructs in a scaffold-free manner. Cultivation of chondrocytes might lead to small cartilage particles, which could be used to replace injured or outworn cartilage. Restoring normal osteogenic differentiation of MSCs from s- exposure by daily short-term stimulation could be helpful so that tissue products may become commercially available, like it has already happened for some bone tissues (BioSeed-Oral Bone, co.don osteotransplant and Osteocel).

DeviceWorking principle

Free fall machineFFMFree fall for 800 ms, “bounce” of 20 g for 50 ms

Levitating magnetsLMA high gradient magnetic field prevents sedimentation

2D-clinostatRotation along one axis

Random positioning machineRPMTwo frames with randomized movement

Rotating wall vesselRWVConstant rotation prevents cells from settling

At the moment, studies analyzing the molecular mechanisms behind spheroid formation of, for example, thyroid cells, chondrocyte, and others have increased the knowledge of the complex regulation of 3D growth in [4245, 171].

To be able to use this new technique more efficiently, further studies are necessary to better understand the exact cellular changes specific to these conditions. Tissue, which was produced under s- or r- conditions, might be helpful to better understand cell signaling, intercellular contact, and tissue growth as well as being sufficient for medical transplantation. MCTS can be used as an alternative to animal experiments.

Although the environment is not a common field for biologists and medical researchers, recent studies have clearly shown that the loss of gravity impacts the cells and it dramatically changes the genome, proteome, and secretome of these cells [43, 45]. Therefore, it is important to systematically explore the advantages of this new research opportunity. Different space flights have already demonstrated a 3D cell growth (Table 2) and similar results have been detected with the help of devices simulating in ground-based laboratories [34, 44, 46, 172174].

Cell typeEngineered tissueMethod References

OsteoblastsBoneRWV bioreactor (s- )[812]
Osteoprogenitor cellsBoneRWV bioreactor (s- )[13, 14]
Mesenchymal stem cellsBoneRWV bioreactor (s- )[12, 1523]
Mesenchymal stem cellsDiversRWV bioreactor (s- )[18, 21, 2426]
Mesenchymal stem cellsDiversRPM (s- )[2730]
Several cell typesMCTSRWV bioreactor (s- )[3134]
Several cell typesMCTSSpinner flask (s- )[3538]
Several cell typesMCTSRPM (s- )[3944]
Several cell typesMCTSSpace (r- )[45]
Hepatocytes liverRWV bioreactor (s- )[34, 46]

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.


This paper was supported by the German Space Agency DLR (Daniela Grimm) (BMWi Grants 50WB0824 and 50WB1124), the European Space Agency (ESA; CORA-GBF-2013-001; CORA-GBF-2013-004), and Aarhus University, Denmark (Daniela Grimm). The authors would like to thank Mr. Peter Lindborg, Los Angeles, CA, USA, for reviewing the paper for language and grammar errors.


  1. R. J. White and M. Averner, “Humans in space,” Nature, vol. 409, no. 6823, pp. 1115–1118, 2001. View at: Publisher Site | Google Scholar
  2. A. I. Grigoriev, V. S. Oganov, A. V. Bakulin et al., “Clinical and physiological evaluation of bone changes among astronauts after long-term space flights,” Aviakosmicheskaia i Ekologicheskaia Meditsina, vol. 32, no. 1, pp. 21–25, 1998. View at: Google Scholar
  3. L. Vico, P. Collet, A. Guignandon et al., “Effects of long-term microgravity exposure on cancellous and cortical weight-bearing bones of cosmonauts,” The Lancet, vol. 355, no. 9215, pp. 1607–1611, 2000. View at: Publisher Site | Google Scholar
  4. S. Tavella, A. Ruggiu, A. Giuliani et al., “Bone turnover in wild type and pleiotrophin-transgenic mice housed for three months in the International Space Station (ISS),” PLoS ONE, vol. 7, no. 3, Article ID e33179, 2012. View at: Publisher Site | Google Scholar
  5. J. Pietsch, J. Bauer, M. Egli et al., “The effects of weightlessness on the human organism and mammalian cells,” Current Molecular Medicine, vol. 11, no. 5, pp. 350–364, 2011. View at: Publisher Site | Google Scholar
  6. D. Grimm, P. Wise, M. Lebert, P. Richter, and S. Baatout, “How and why does the proteome respond to microgravity?” Expert Review of Proteomics, vol. 8, no. 1, pp. 13–27, 2011. View at: Publisher Site | Google Scholar
  7. D. Grimm, M. Wehland, J. Pietsch J et al., “Growing tissues in real and simulated microgravity: new methods for tissue engineering,” Tissue Engineering Part B Reviews, 2014. View at: Publisher Site | Google Scholar
  8. C. Granet, N. Laroche, L. Vico, C. Alexandre, and M. H. Lafage-Proust, “Rotating-wall vessels, promising bioreactors for osteoblastic cell culture: comparison with other 3D conditions,” Medical and Biological Engineering and Computing, vol. 36, no. 4, pp. 513–519, 1998. View at: Publisher Site | Google Scholar
  9. K. Song, Z. Yang, T. Liu et al., “Fabrication and detection of tissue-engineered bones with bio-derived scaffolds in a rotating bioreactor,” Biotechnology and Applied Biochemistry, vol. 45, no. 2, pp. 65–74, 2006. View at: Publisher Site | Google Scholar
  10. K. Song, T. Liu, Z. Cui, X. Li, and X. Ma, “Three-dimensional fabrication of engineered bone with human bio-derived bone scaffolds in a rotating wall vessel bioreactor,” Journal of Biomedical Materials Research A, vol. 86, no. 2, pp. 323–332, 2008. View at: Publisher Site | Google Scholar
  11. K. Kyriakidou, G. Lucarini, A. Zizzi et al., “Dynamic co-seeding of osteoblast and endothelial cells on 3D polycaprolactone scaffolds for enhanced bone tissue engineering,” Journal of Bioactive and Compatible Polymers, vol. 23, no. 3, pp. 227–243, 2008. View at: Publisher Site | Google Scholar
  12. Q. Lv, L. Nair, and C. T. Laurencin, “Fabrication, characterization, and in vitro evaluation of poly(lactic acid glycolic acid)/nano-hydroxyapatite composite microsphere-based scaffolds for bone tissue engineering in rotating bioreactors,” Journal of Biomedical Materials Research A, vol. 91, no. 3, pp. 679–691, 2009. View at: Publisher Site | Google Scholar
  13. S. R. Facer, R. S. Zaharias, M. E. Andracki, J. Lafoon, S. K. Hunter, and G. B. Schneider, “Rotary culture enhances pre-osteoblast aggregation and mineralization,” Journal of Dental Research, vol. 84, no. 6, pp. 542–547, 2005. View at: Publisher Site | Google Scholar
  14. G. B. Schneider, J. K. Boehrs, J. V. Hoopes, and D. A. Seabold, “Use of 3-dimensional environments to engineer osseous-like tissue,” Journal of Developmental Biology and Tissue Engineering, vol. 3, no. 4, pp. 42–47, 2011. View at: Google Scholar
  15. D. Turhani, E. Watzinger, M. Weißenböck et al., “Analysis of cell-seeded 3-dimensional bone constructs manufactured in vitro with hydroxyapatite granules obtained from red algae,” Journal of Oral and Maxillofacial Surgery, vol. 63, no. 5, pp. 673–681, 2005. View at: Publisher Site | Google Scholar
  16. F. Jin, Y. Zhang, K. Xuan et al., “Establishment of three-dimensional tissue-engineered bone constructs under microgravity-simulated conditions,” Artificial Organs, vol. 34, no. 2, pp. 118–125, 2010. View at: Publisher Site | Google Scholar
  17. J. G. Gershovich and L. B. Buravkova, “Morphofunctional status and osteogenic differentiation potential of human mesenchymal stromal precursor cells during in vitro modeling of microgravity effects,” Bulletin of Experimental Biology and Medicine, vol. 144, no. 4, pp. 608–613, 2007. View at: Publisher Site | Google Scholar
  18. M. Zayzafoon, W. E. Gathings, and J. M. McDonald, “Modeled microgravity inhibits osteogenic differentiation of human mesenchymal stem cells and increases adipogenesis,” Endocrinology, vol. 145, no. 5, pp. 2421–2432, 2004. View at: Publisher Site | Google Scholar
  19. R. Saxena, G. Pan, and J. M. McDonald, “Osteoblast and osteoclast differentiation in modeled microgravity,” Annals of the New York Academy of Sciences, vol. 1116, pp. 494–498, 2007. View at: Publisher Site | Google Scholar
  20. V. E. Meyers, M. Zayzafoon, S. R. Gonda, W. E. Gathings, and J. M. McDonald, “Modeled microgravity disrupts collagen I/integrin signaling during osteoblastic differentiation of human mesenchymal stem cells,” Journal of Cellular Biochemistry, vol. 93, no. 4, pp. 697–707, 2004. View at: Publisher Site | Google Scholar
  21. D. Sheyn, G. Pelled, D. Netanely, E. Domany, and D. Gazit, “The effect of simulated microgravity on human mesenchymal stem cells cultured in an osteogenic differentiation system: a bioinformatics study,” Tissue Engineering A, vol. 16, no. 11, pp. 3403–3412, 2010. View at: Publisher Site | Google Scholar
  22. B. Yu, D. Yu, L. Cao et al., “Simulated microgravity using a rotary cell culture system promotes chondrogenesis of human adipose-derived mesenchymal stem cells via the p38 MAPK pathway,” Biochemical and Biophysical Research Communications, vol. 414, no. 2, pp. 412–418, 2011. View at: Publisher Site | Google Scholar
  23. S. M. Uddin and Y. X. Qin, “Enhancement of osteogenic differentiation and proliferation in human mesenchymal stem cells by a modified low intensity ultrasound stimulation under simulated microgravity,” PLoS ONE, vol. 8, no. 9, Article ID e73914, 2013. View at: Google Scholar
  24. Z. Q. Dai, R. Wang, S. K. Ling, Y. M. Wan, and Y. H. Li, “Simulated microgravity inhibits the proliferation and osteogenesis of rat bone marrow mesenchymal stem cells,” Cell Proliferation, vol. 40, no. 5, pp. 671–684, 2007. View at: Publisher Site | Google Scholar
  25. X. Wu, S.-H. Li, L.-M. Lou, and Z.-R. Chen, “The effect of the microgravity rotating culture system on the chondrogenic differentiation of bone marrow mesenchymal stem cells,” Molecular Biotechnology, vol. 54, no. 2, pp. 331–336, 2013. View at: Publisher Site | Google Scholar
  26. X. Zhang, Y. Nan, H. Wang et al., “Model microgravity enhances endothelium differentiation of mesenchymal stem cells,” Naturwissenschaften, vol. 100, no. 2, pp. 125–133, 2013. View at: Publisher Site | Google Scholar
  27. N. V. Merzlikina, L. B. Buravkova, and Y. A. Romanov, “The primary effects of clinorotation on cultured human mesenchymal stem cells,” Journal of Gravitational Physiology, vol. 11, no. 2, pp. 193–194, 2004. View at: Google Scholar
  28. M. Nishikawa, H. Ohgushi, N. Tamai et al., “The effect of simulated microgravity by three-dimensional clinostat on bone tissue engineering,” Cell Transplantation, vol. 14, no. 10, pp. 829–835, 2005. View at: Publisher Site | Google Scholar
  29. L. Yuge, T. Kajiume, H. Tahara et al., “Microgravity potentiates stem cell proliferation while sustaining the capability of differentiation,” Stem Cells and Development, vol. 15, no. 6, pp. 921–929, 2006. View at: Publisher Site | Google Scholar
  30. I. G. Gershovich and L. B. Buravkova, “Interleukine production in culture of mesenchymal stromal cells of humans during simulation of the microgravity effects,” Aviakosmicheskaia I Ekologicheskaia Meditsina, vol. 43, no. 3, pp. 44–50, 2009. View at: Google Scholar
  31. M. Ingram, G. B. Techy, R. Saroufeem et al., “Three-dimensional growth patterns of various human tumor cell lines in simulated microgravity of a nasa bioreactor,” In Vitro Cellular and Developmental Biology: Animal, vol. 33, no. 6, pp. 459–466, 1997. View at: Publisher Site | Google Scholar
  32. B. Marrero, J. L. Messina, and R. Heller, “Generation of a tumor spheroid in a microgravity environment as a 3D model of melanoma,” In Vitro Cellular and Developmental Biology—Animal, vol. 45, no. 9, pp. 523–534, 2009. View at: Publisher Site | Google Scholar
  33. S. M. Konstantinov, M. M. Mindova, P. T. Gospodinov, and P. I. Genova, “Three-dimensional bioreactor cultures: a useful dynamic model for the study of cellular interactions,” Annals of the New York Academy of Sciences, vol. 1030, pp. 103–115, 2004. View at: Publisher Site | Google Scholar
  34. T. T. Chang and M. Hughes-Fulford, “Monolayer and spheroid culture of human liver hepatocellular carcinoma cell line cells demonstrate distinct global gene expression patterns and functional phenotypes,” Tissue Engineering A, vol. 15, no. 3, pp. 559–567, 2009. View at: Publisher Site | Google Scholar
  35. A. Bredel-Geissler, U. Karbach, S. Walenta, L. Vollrath, and W. Mueller-Klieser, “Proliferation-associated oxygen consumption and morphology of tumor cells in monolayer and spheroid culture,” Journal of Cellular Physiology, vol. 153, no. 1, pp. 44–52, 1992. View at: Publisher Site | Google Scholar
  36. L. A. Kunz-Schughart, C. Groebe, and W. Mueller-Klieser, “Three-dimensional cell culture induces novel proliferative and metabolic alterations associated with oncogenic transformation,” International Journal of Cancer, vol. 66, no. 4, pp. 578–586, 1996. View at: Google Scholar
  37. M. Wartenberg, F. Dönmez, F. C. Ling, H. Acker, J. Hescheler, and H. Sauer, “Tumor-induced angiogenesis studied in confrontation cultures of multicellular tumor spheroids and embryoid bodies grown from pluripotent embryonic stem cells,” The FASEB Journal, vol. 15, no. 6, pp. 995–1005, 2001. View at: Publisher Site | Google Scholar
  38. R.-Z. Lin and H.-Y. Chang, “Recent advances in three-dimensional multicellular spheroid culture for biomedical research,” Biotechnology Journal, vol. 3, no. 9-10, pp. 1172–1184, 2008. View at: Publisher Site | Google Scholar
  39. D. Grimm, J. Bauer, P. Kossmehl et al., “Simulated microgravity alters differentiation and increases apoptosis in human follicular thyroid carcinoma cells,” The FASEB Journal, vol. 16, no. 6, pp. 604–606, 2002. View at: Google Scholar
  40. M. Infanger, P. Kossmehl, M. Shakibaei et al., “Simulated weightlessness changes the cytoskeleton and extracellular matrix proteins in papillary thyroid carcinoma cells,” Cell and Tissue Research, vol. 324, no. 2, pp. 267–277, 2006. View at: Publisher Site | Google Scholar
  41. J. Grosse, M. Wehland, J. Pietsch et al., “Gravity-sensitive signaling drives 3-dimensional formation of multicellular thyroid cancer spheroids,” The FASEB Journal, vol. 26, no. 12, pp. 5124–5140, 2012. View at: Publisher Site | Google Scholar
  42. X. Ma, A. Sickmann, J. Pietsch et al., “Proteomic differences between microvascular endothelial cells and the EA.hy926 cell line forming three-dimensional structures,” Proteomics, vol. 14, no. 6, pp. 689–698, 2014. View at: Google Scholar
  43. X. Ma, J. Pietsch, M. Wehland et al., “Differential gene expression profile and altered cytokine secretion of thyroid cancer cells in space,” FASEB Journal, vol. 28, no. 2, pp. 813–835, 2014. View at: Publisher Site | Google Scholar
  44. J. Pietsch, A. Sickmann, G. Weber et al., “A proteomic approach to analysing spheroid formation of two human thyroid cell lines cultured on a random positioning machine,” Proteomics, vol. 11, no. 10, pp. 2095–2104, 2011. View at: Publisher Site | Google Scholar
  45. J. Pietsch, X. Ma, M. Wehland et al., “Spheroid formation of human thyroid cancer cells in an automated culturing system during the Shenzhou-8 Space mission,” Biomaterials, vol. 34, no. 31, pp. 7694–7705, 2013. View at: Publisher Site | Google Scholar
  46. T. T. Chang and M. Hughes-Fulford, “Molecular mechanisms underlying the enhanced functions of three-dimensional hepatocyte aggregates,” Biomaterials, vol. 35, no. 7, pp. 2162–2171, 2014. View at: Google Scholar
  47. M. Y. Kapitonova, N. Salim, S. Othman et al., “Alteration of cell cytoskeleton and functions of cell recovery of normal human osteoblast cells caused by factors associated with real space flight,” The Malaysian Journal of Pathology, vol. 35, no. 2, pp. 153–163, 2013. View at: Google Scholar
  48. E. A. Blaber, N. Dvorochkin, C. Lee et al., “Microgravityinduces pelvic bone loss through osteoclastic activity, osteocytic osteolysis, and osteoblastic cell cycle inhibition by CDKN1α/p21,” PLoS ONE, vol. 8, no. 4, Article ID 61372, 2013. View at: Publisher Site | Google Scholar
  49. C. Ulbrich, J. Pietsch, J. Grosse et al., “Differential gene regulation under altered gravity conditions in follicular thyroid cancer cells: relationship between the extracellular matrix and the cytoskeleton,” Cellular Physiology and Biochemistry, vol. 28, no. 2, pp. 185–198, 2011. View at: Publisher Site | Google Scholar
  50. J. Grosse, M. Wehland, J. Pietsch et al., “Short-term weightlessness produced by parabolic flight maneuvers altered gene expression patterns in human endothelial cells,” The FASEB Journal, vol. 26, no. 2, pp. 639–655, 2012. View at: Publisher Site | Google Scholar
  51. R. Hemmersbach, M. Krause, R. Bräucker, and K. Ivanova, “Graviperception in ciliates: steps in the transduction chain,” Advances in Space Research, vol. 35, no. 2, pp. 296–299, 2005. View at: Publisher Site | Google Scholar
  52. D. A. M. Mesland, “Novel ground-based facilities for research in the effects of weight,” ESA Microgravity News, vol. 9, 1996. View at: Google Scholar
  53. D. A. M. Meslana, A. H. Anton, H. Willemsen, and H. van den Ende, “The free fall machine—a ground-based facility for microgravity research in life sciences,” Microgravity Science and Technology, vol. 9, no. 1, pp. 10–14, 1996. View at: Google Scholar
  54. M. Schwarzenberg, P. Pippia, M. A. Meloni, G. Cossu, M. Cogoli-Greuter, and A. Cogoli, “Signal transduction in T lymphocytes—a comparison of the data from space, the free fall machine and the random positioning machine,” Advances in Space Research, vol. 24, no. 6, pp. 793–800, 1999. View at: Publisher Site | Google Scholar
  55. M. V. Berry and A. K. Geim, “Of flying frogs and levitrons,” European Journal of Physics, vol. 18, no. 4, pp. 307–313, 1997. View at: Publisher Site | Google Scholar | MathSciNet
  56. J. M. Valles Jr., K. Lin, J. M. Denegre, and K. L. Mowry, “Stable magnetic field gradient levitation of Xenopus laevis: toward low- gravity simulation,” Biophysical Journal, vol. 73, no. 2, pp. 1130–1133, 1997. View at: Publisher Site | Google Scholar
  57. M. J. A. Moes, J. C. Gielen, R.-J. Bleichrodt, J. J. W. A. van Loon, P. C. M. Christianen, and J. Boonstra, “Simulation of microgravity by magnetic levitation and random positioning: effect on human A431 Cell morphology,” Microgravity Science and Technology, vol. 23, no. 2, pp. 249–261, 2011. View at: Publisher Site | Google Scholar
  58. R. Herranz, R. Anken, J. Boonstra et al., “Ground-based facilities for simulation of microgravity: organism-specific recommendations for their use, and recommended terminology,” Astrobiology, vol. 13, no. 1, pp. 1–17, 2013. View at: Publisher Site | Google Scholar
  59. T. Higashi, A. Yamagishi, T. Takeuchi et al., “Orientation of erythrocytes in a strong static magnetic field,” Blood, vol. 82, no. 4, pp. 1328–1334, 1993. View at: Google Scholar
  60. K. A. Mirica, F. Ilievski, A. K. Ellerbee, S. S. Shevkoplyas, and G. M. Whitesides, “Using magnetic levitation for three dimensional self-assembly,” Advanced Materials, vol. 23, no. 36, pp. 4134–4140, 2011. View at: Publisher Site | Google Scholar
  61. J. S. Brooks, J. A. Reavis, R. A. Medwood et al., “New opportunities in science, materials, and biological systems in the low-gravity (magnetic levitation) environment (invited),” Journal of Applied Physics, vol. 87, no. 9, pp. 6194–6199, 2000. View at: Publisher Site | Google Scholar
  62. B. E. Hammer, L. S. Kidder, P. C. Williams, and W. W. Xu, “Magnetic levitation of MC3T3 osteoblast cells as a ground-based simulation of microgravity,” Microgravity Science and Technology, vol. 21, no. 4, pp. 311–318, 2009. View at: Publisher Site | Google Scholar
  63. C. E. Dijkstra, O. J. Larkin, P. Anthony et al., “Diamagnetic levitation enhances growth of liquid bacterial cultures by increasing oxygen availability,” Journal of the Royal Society Interface, vol. 8, no. 56, pp. 334–344, 2011. View at: Publisher Site | Google Scholar
  64. D. Shi, R. Meng, W. Deng et al., “Effects of microgravity modeled by large gradient high magnetic field on the osteogenic initiation of human mesenchymal stem cells,” Stem Cell Reviews and Reports, vol. 6, no. 4, pp. 567–578, 2010. View at: Publisher Site | Google Scholar
  65. G. R. Souza, J. R. Molina, R. M. Raphael et al., “Three-dimensional tissue culture based on magnetic cell levitation,” Nature Nanotechnology, vol. 5, no. 4, pp. 291–296, 2010. View at: Publisher Site | Google Scholar
  66. F. Castro-Chavez, K. C. Vickers, J. S. Lee, C. Tung, and J. D. Morrisett, “Effect of lyso-phosphatidylcholine and Schnurri-3 on osteogenic transdifferentiation of vascular smooth muscle cells to calcifying vascular cells in 3D culture,” Biochimica et Biophysica Acta, vol. 1830, no. 6, pp. 3828–3834, 2013. View at: Publisher Site | Google Scholar
  67. F. G. J. R. von Sachs, “Über Ausschliessung der geotropischen und heliotropischen Krümmungen während des Wachstums,” Würzburger Arbeiten, vol. 2, pp. 209–225, 1879. View at: Google Scholar
  68. J. J. W. A. van Loon, “Some history and use of the random positioning machine, RPM, in gravity related research,” Advances in Space Research, vol. 39, no. 7, pp. 1161–1165, 2007. View at: Publisher Site | Google Scholar
  69. S. Murakami and M. Yamada, “Architecture of statocytes and chloroplasts under the microgravity environment,” Biological Sciences in Space, vol. 2, no. 4, p. 301, 1988. View at: Google Scholar
  70. J. J. W. A. van Loon, “Effects of spaceflight on biological systems,” in Biology in Space and Life on Earth, E. Brinckmann, Ed., vol. 17, Wiley-VCH, 2007. View at: Google Scholar
  71. T. Hoson, S. Kamisaka, Y. Masuda, and M. Yamashita, “Changes in plant growth processes under microgravity conditions simulated by a three-dimensional clinostat,” The Botanical Magazine Tokyo, vol. 105, no. 1, pp. 53–70, 1992. View at: Publisher Site | Google Scholar
  72. T. Hoson, S. Kamisaka, Y. Masuda, M. Yamashita, and B. Buchen, “Evaluation of the three-dimensional clinostat as a simulator of weightlessness,” Planta, vol. 203, pp. S187–S197, 1997. View at: Publisher Site | Google Scholar
  73. A. G. Borst and J. J. W. A. van Loon, “Technology and developments for the random positioning machine, RPM,” Microgravity Science and Technology, vol. 21, no. 4, pp. 287–292, 2009. View at: Publisher Site | Google Scholar
  74. C. A. D. Leguy, R. Delfos, and M. J. B. M Pourquie, “Fluid motion for microgravity simulations in a random positioning machine,” Gravitational and Space Biology, vol. 25, no. 1, pp. 36–39, 2011. View at: Google Scholar
  75. S. Wakayama, Y. Kawahara, C. Li, K. Yamagata, L. Yuge, and T. Wakayama, “Detrimental effects of microgravity on mouse preimplantation development in vitro,” PLoS ONE, vol. 4, no. 8, Article ID e6753, 2009. View at: Publisher Site | Google Scholar
  76. M. Z. Luo, R. Meng, S. S. Li et al., “Weightlessness simulated with random positioning machine influences the cytoskeleton and migration of MC3T3-E1 cells,” Journal of the Japan Society of Microgravity Application, vol. 28, no. 2, pp. S41–S45, 2011. View at: Google Scholar
  77. W. A. Loesberg, X. F. Walboomers, E. M. Bronkhorst, J. J. W. A. Van Loon, and J. A. Jansen, “The effect of combined simulated microgravity and microgrooved surface topography on fibroblasts,” Cell Motility and the Cytoskeleton, vol. 64, no. 3, pp. 174–185, 2007. View at: Publisher Site | Google Scholar
  78. G. Aleshcheva, J. Sahana, X. Ma et al., “Changes in morphology, gene expression and protein content in chondrocytes cultured on a Random Positioning Machine,” PLoS ONE, vol. 8, no. 11, Article ID e79057, 2013. View at: Google Scholar
  79. L. Prodanov, J. J. W. A. van Loon, J. te Riet, J. A. Jansen, and X. F. Walboomers, “Nanostructured substrate conformation can decrease osteoblast-like cell dysfunction in simulated microgravity conditions,” Journal of Tissue Engineering and Regenerative Medicine, 2012. View at: Publisher Site | Google Scholar
  80. J. Ichigi and M. Asashima, “Dome formation and tubule morphogenesis by Xenopus kidney A6 cell cultures exposed to microgravity simulated with a 3D-clinostat and to hypergravity,” In Vitro Cellular Developmental Biology. Animal, vol. 37, no. 1, pp. 31–44, 2001. View at: Publisher Site | Google Scholar
  81. Y. D. Jiang, W. N. Li, L. F. Wang, Z. Y. Zhang, B. M. Zhang, and H. J. Wu, “Several new types of clinostats,” Space Medicine & Medical Engineering, vol. 21, no. 4, pp. 368–371, 2008. View at: Google Scholar
  82. J. H. Siamwala, S. H. Reddy, S. Majumder et al., “Simulated microgravity perturbs actin polymerization to promote nitric oxide-associated migration in human immortalized Eahy926 cells,” Protoplasma, vol. 242, no. 1, pp. 3–12, 2010. View at: Publisher Site | Google Scholar
  83. L. Xiang, F. Qi, D. Dai, C. Li, and Y. Jiang, “Simulated microgravity affects growth of Escherichia coli and recombinant β-D-glucuronidase production,” Applied Biochemistry and Biotechnology, vol. 162, no. 3, pp. 654–661, 2010. View at: Publisher Site | Google Scholar
  84. R. Hemmersbach, S. M. Strauch, D. Seibt, and M. Schuber, “Comparative studies on gravisensitive protists on ground (2D and 3D clinostats) and in microgravity,” Microgravity Science and Technology, vol. 18, no. 3-4, pp. 257–259, 2006. View at: Publisher Site | Google Scholar
  85. T. J. Goodwin, J. M. Jessup, and D. A. Wolf, “Morphologic differentiation of colon carcinoma cell lines HT-29 and HT-29 KM in Rotating-Wall Vessels,” In Vitro Cellular and Developmental Biology: Animal, vol. 28, no. 1, pp. 47–60, 1992. View at: Publisher Site | Google Scholar
  86. T. G. Hammond and J. M. Hammond, “Optimized suspension culture: the rotating-wall vessel,” The American Journal of Physiology—Renal Physiology, vol. 281, no. 1, pp. F12–F25, 2001. View at: Google Scholar
  87. T. Liu, X. Li, X. Sun, X. Ma, and Z. Cui, “Analysis on forces and movement of cultivated particles in a rotating wall vessel bioreactor,” Biochemical Engineering Journal, vol. 18, no. 2, pp. 97–104, 2004. View at: Publisher Site | Google Scholar
  88. T. J. Goodwin, T. L. Prewett, D. A. Wolf, and G. F. Spaulding, “Reduced shear stress: a major component in the ability of mammalian tissues to form three-dimensional assemblies in simulated microgravity,” Journal of Cellular Biochemistry, vol. 51, no. 3, pp. 301–311, 1993. View at: Publisher Site | Google Scholar
  89. E. A. Nauman, C. M. Ott, E. Sander et al., “Novel quantitative biosystem for modeling physiological fluid shear stress on cells,” Applied and Environmental Microbiology, vol. 73, no. 3, pp. 699–705, 2007. View at: Publisher Site | Google Scholar
  90. X. Guo, Y. Zhao, H. Chang et al., “Creation of engineered cardiac tissue in vitro from mouse embryonic stem cells,” Circulation, vol. 113, no. 18, pp. 2229–2237, 2006. View at: Publisher Site | Google Scholar
  91. D. L. Tucker, C. M. Ott, S. Huff et al., “Characterization of Escherichia coli MG1655 grown in a low-shear modeled microgravity environment,” BMC Microbiology, vol. 7, article 15, 2007. View at: Publisher Site | Google Scholar
  92. M. N. Cinbiz, R. S. Tıgli, I. G. Beşkardeş, M. Gümüşderelioğlu, and U. Colak, “Computational fluid dynamics modeling of momentum transport in rotating wall perfused bioreactor for cartilage tissue engineering,” Journal of Biotechnology, vol. 150, no. 3, pp. 389–395, 2010. View at: Google Scholar
  93. A. Puca, G. Russo, and A. Giordano, “Properties of mechano-transduction via simulated microgravity and its effects on intracellular trafficking of VEGFR's,” Oncotarget, vol. 3, no. 4, pp. 426–434, 2012. View at: Google Scholar
  94. J. H. Siamwala, S. Majumder, K. P. Tamilarasan et al., “Simulated microgravity promotes nitric oxide-supported angiogenesis via the iNOS-cGMP-PKG pathway in macrovascular endothelial cells,” FEBS Letters, vol. 584, no. 15, pp. 3415–3423, 2010. View at: Publisher Site | Google Scholar
  95. Y. C. Wang, S. Zhang, T. Y. Du, B. Wang, and X. Q. Sun, “Clinorotation upregulates inducible nitric oxide synthase by inhibiting AP-1 activation in human umbilical vein endothelial cells,” Journal of Cellular Biochemistry, vol. 107, no. 2, pp. 357–363, 2009. View at: Publisher Site | Google Scholar
  96. F. Shi, Y.-C. Wang, T.-Z. Zhao et al., “Effects of simulated microgravity on human umbilical vein endothelial cell angiogenesis and role of the PI3K-Akt-eNOS signal pathway,” PLoS ONE, vol. 7, no. 7, Article ID e40365, 2012. View at: Publisher Site | Google Scholar
  97. P. I. Lelkes, N. Akhtar, E. Lelkes et al., “Neuroendocrine tissue engineering in rotating wall vessel bioreactors under simulated microgravity conditions,” in Proceedings of the 23rd Annual International Conference of the IEEE Engineering in Medicine and Biology Society, vol. 3, pp. 2987–2990, October 2001. View at: Publisher Site | Google Scholar
  98. A. Abbott, “Cell culture: biology’s new dimensions,” Nature, vol. 424, no. 6951, pp. 870–872, 2003. View at: Publisher Site | Google Scholar
  99. B. R. Unsworth and P. I. Lelkes, “Growing tissues in microgravity,” Nature Medicine, vol. 4, no. 8, pp. 901–907, 1998. View at: Publisher Site | Google Scholar
  100. M. Infanger, P. Kossmehl, M. Shakibaei et al., “Induction of three-dimensional assembly and increase in apoptosis of human endothelial cells by simulated microgravity: impact of vascular endothelial growth factor,” Apoptosis, vol. 11, no. 5, pp. 749–764, 2006. View at: Publisher Site | Google Scholar
  101. D. Grimm, M. Infanger, K. Westphal et al., “A delayed type of three-dimensional growth of human endothelial cells under simulated weightlessness,” Tissue Engineering A, vol. 15, no. 8, pp. 2267–2275, 2009. View at: Publisher Site | Google Scholar
  102. D. Grimm, J. Bauer, C. Ulbrich et al., “Different responsiveness of endothelial cells to vascular endothelial growth factor and basic fibroblast growth factor added to culture media under gravity and simulated microgravity,” Tissue Engineering A, vol. 16, no. 5, pp. 1559–1573, 2010. View at: Publisher Site | Google Scholar
  103. M. Infanger, C. Ulbrich, S. Baatout et al., “Modeled gravitational unloading induced downregulation of endothelin-1 in human endothelial cells,” Journal of Cellular Biochemistry, vol. 101, no. 6, pp. 1439–1455, 2007. View at: Publisher Site | Google Scholar
  104. X. Ma, M. Wehland, H. Schulz et al., “Genomic Approach to identify factors that drive the formation of three-dimensional structures by EA.hy926 endothelial cells,” PLoS ONE, vol. 8, no. 5, Article ID e64402, 2013. View at: Publisher Site | Google Scholar
  105. C. Ulbrich, K. Westphal, J. Pietsch et al., “Characterization of human chondrocytes exposed to simulated microgravity,” Cellular Physiology and Biochemistry, vol. 25, no. 4-5, pp. 551–560, 2010. View at: Publisher Site | Google Scholar
  106. L. E. Freed and G. Vunjak-Novakovic, “Cultivation of cell-polymer tissue constructs in simulated microgravity,” Biotechnology and Bioengineering, vol. 46, no. 4, pp. 306–313, 1995. View at: Publisher Site | Google Scholar
  107. L. E. Freed, R. Langer, I. Martin, N. R. Pellis, and G. Vunjak-Novakovic, “Tissue engineering of cartilage in space,” Proceedings of the National Academy of Sciences of the United States of America, vol. 94, no. 25, pp. 13885–13890, 1997. View at: Publisher Site | Google Scholar
  108. V. Stamenković, G. Keller, D. Nesic, A. Cogoli, and S. P. Grogan, “Neocartilage formation in 1 g, simulated, and microgravity environments: implications for tissue engineering,” Tissue Engineering A, vol. 16, no. 5, pp. 1729–1736, 2010. View at: Publisher Site | Google Scholar
  109. L. Kock, C. C. van Donkelaar, and K. Ito, “Tissue engineering of functional articular cartilage: the current status,” Cell and Tissue Research, vol. 347, no. 3, pp. 613–627, 2012. View at: Publisher Site | Google Scholar
  110. C. H. Chang, F. H. Lin, T. F. Kuo, and H. C. Liu, “Cartilage tissue engineering,” Biomedical Engineering: Applications, Basis and Communications, vol. 17, no. 2, pp. 61–71, 2005. View at: Publisher Site | Google Scholar
  111. E. Ozcivici, Y. K. Luu, B. Adler et al., “Mechanical signals as anabolic agents in bone,” Nature Reviews Rheumatology, vol. 6, no. 1, pp. 50–59, 2010. View at: Publisher Site | Google Scholar
  112. R. Yang, W. Lin, Y. Chen et al., “Regulation by ultrasound treatment on the integrin expression and differentiation of osteoblasts,” Bone, vol. 36, no. 2, pp. 276–283, 2005. View at: Publisher Site | Google Scholar
  113. H. C. Hsu, Y. C. Fong, C. S. Chang et al., “Ultrasound induces cyclooxygenase-2 expression through integrin, integrin-linked kinase, Akt, NF-κB and p300 pathway in human chondrocytes,” Cellular Signalling, vol. 19, no. 11, pp. 2317–2328, 2007. View at: Publisher Site | Google Scholar
  114. Y. I. Arfat, W. Z. Xiao, S. Iftikhar et al., “Physiological effects of microgravity on bone cells,” Calcified Tissue International, vol. 94, no. 6, pp. 569–579, 2014. View at: Publisher Site | Google Scholar
  115. D. A. Starr, “Communication between the cytoskeleton and the nuclear envelope to position the nucleus,” Molecular BioSystems, vol. 3, no. 9, pp. 583–589, 2007. View at: Publisher Site | Google Scholar
  116. N. Nabavi, A. Khandani, A. Camirand, and R. E. Harrison, “Effects of microgravity on osteoclast bone resorption and osteoblast cytoskeletal organization and adhesion,” Bone, vol. 49, no. 5, pp. 965–974, 2011. View at: Publisher Site | Google Scholar
  117. S. B. Doty, E. R. Morey-Holton, G. N. Durnova, and A. S. Kaplansky, “Cosmos 1887: morphology, histochemistry, and vasculature of the growing rat tibia,” FASEB Journal, vol. 4, no. 1, pp. 16–23, 1990. View at: Google Scholar
  118. N. V. Rodionova, O. V. Polkovenko, and V. S. Oganov, “Interactions of cells in zones of bone resorption under microgravity and hypokinesia.,” Journal of Gravitational Physiology, vol. 11, no. 2, pp. P147–151, 2004. View at: Google Scholar
  119. O. D. Kennedy, B. C. Herman, D. M. Laudier, R. J. Majeska, H. B. Sun, and M. B. Schaffler, “Activation of resorption in fatigue-loaded bone involves both apoptosis and active pro-osteoclastogenic signaling by distinct osteocyte populations,” Bone, vol. 50, no. 5, pp. 1115–1122, 2012. View at: Publisher Site | Google Scholar
  120. C. Lin, X. Jiang, Z. Dai et al., “Sclerostin mediates bone response to mechanical unloading through antagonizing Wnt/β-catenin signaling,” Journal of Bone and Mineral Research, vol. 24, no. 10, pp. 1651–1661, 2009. View at: Publisher Site | Google Scholar
  121. R. Tamma, G. Colaianni, C. Camerino et al., “Microgravity during spaceflight directly affects in vitro osteoclastogenesis and bone resorption,” The FASEB Journal, vol. 23, no. 8, pp. 2549–2554, 2009. View at: Publisher Site | Google Scholar
  122. A. Sundaresan, S. F. Clarke, and N. R. Pellis, Production of Bone Morphogenic Proteins (BMPS) Using a Novel Tissue Culture Platform, 2011, US Patent 8076136.
  123. A. Sundaresan, S. F. Clarke, and N. R. Pellis, “Constructions osseuses minéralisées en trois dimensions,” European Patent Number: EP2013730, 2011. View at: Google Scholar
  124. A. Sundaresan, M. S. F. Clarke, and M. Brinker, Development of a human colloidal bone graft material, US 8506982 B2, 2013.
  125. J. R. Milstead, S. J. Simske, and T. A. Bateman, “Spaceflight and hindlimb suspension disuse models in mice,” Biomedical Sciences Instrumentation, vol. 40, pp. 105–110, 2004. View at: Google Scholar
  126. H. M. Frost and W. S. Jee, “On the rat model of human osteopenias and osteoporoses,” Bone and Mineral, vol. 18, no. 3, pp. 227–236, 1992. View at: Publisher Site | Google Scholar
  127. J. H. Keyak, A. K. Koyama, A. LeBlanc, Y. Lu, and T. F. Lang, “Reduction in proximal femoral strength due to long-duration spaceflight,” Bone, vol. 44, no. 3, pp. 449–453, 2009. View at: Publisher Site | Google Scholar
  128. S. M. Smith, M. E. Wastney, K. O. O'Brien et al., “Bone markers, calcium metabolism, and calcium kinetics during extended-duration space flight on the Mir Space Station,” Journal of Bone and Mineral Research, vol. 20, no. 2, pp. 208–218, 2005. View at: Publisher Site | Google Scholar
  129. J. M. Vogel, “Bone mineral measurement: Skylab experiment M-078,” Acta Astronautica, vol. 2, no. 1-2, pp. 129–139, 1975. View at: Publisher Site | Google Scholar
  130. S. M. Smith, M. E. Wastney, B. V. Morukov et al., “Calcium metabolism before, during, and after a 3-mo spaceflight: kinetic and biochemical changes,” American Journal of Physiology, vol. 277, no. 1, part 2, pp. R1–R10, 1999. View at: Google Scholar
  131. J. E. Zerwekh, L. A. Ruml, F. Gottschalk, and C. Y. C. Pak, “The effects of twelve weeks of bed rest on bone histology, biochemical markers of bone turnover, and calcium homeostasis in eleven normal subjects,” Journal of Bone and Mineral Research, vol. 13, no. 10, pp. 1594–1601, 1998. View at: Publisher Site | Google Scholar
  132. T. Lang, A. LeBlanc, H. Evans, Y. Lu, H. Genant, and A. Yu, “Cortical and trabecular bone mineral loss from the spine and hip in long-duration spaceflight,” Journal of Bone and Mineral Research, vol. 19, no. 6, pp. 1006–1012, 2004. View at: Publisher Site | Google Scholar
  133. A. LeBlanc, C. Lin, L. Shackelford et al., “Muscle volume, MRI relaxation times (T2), and body composition after spaceflight,” Journal of Applied Physiology, vol. 89, no. 6, pp. 2158–2164, 2000. View at: Google Scholar
  134. M. Iki, E. Kajita, Y. Dohi et al., “Age, menopause, bone turnover markers and lumbar bone loss in healthy Japanese women,” Maturitas, vol. 25, no. 1, pp. 59–67, 1996. View at: Publisher Site | Google Scholar
  135. J. Sirola, H. Kröger, R. Honkanen et al., “Factors affecting bone loss around menopause in women without HRT: a prospective study,” Maturitas, vol. 45, no. 3, pp. 159–167, 2003. View at: Publisher Site | Google Scholar
  136. J. D. Sibonga, H. J. Evans, H. G. Sung et al., “Recovery of spaceflight-induced bone loss: bone mineral density after long-duration missions as fitted with an exponential function,” Bone, vol. 41, no. 6, pp. 973–978, 2007. View at: Publisher Site | Google Scholar
  137. T. F. Lang, A. D. Leblanc, H. J. Evans, and Y. Lu, “Adaptation of the proximal femur to skeletal reloading after long-duration spaceflight,” Journal of Bone and Mineral Research, vol. 21, no. 8, pp. 1224–1230, 2006. View at: Publisher Site | Google Scholar
  138. J. J. W. A. Van Loon, D.-J. Bervoets, E. H. Burger et al., “Decreased mineralization and increased calcium release in isolated fetal mouse long bones under near weightlessness,” Journal of Bone and Mineral Research, vol. 10, no. 4, pp. 550–557, 1995. View at: Google Scholar
  139. R. Langer, “Tissue engineering: a new field and its challenges,” Pharmaceutical Research, vol. 14, no. 7, pp. 840–841, 1997. View at: Publisher Site | Google Scholar
  140. G. M. Crane, S. L. Ishaug, and A. G. Mikos, “Bone tissue engineering,” Nature Medicine, vol. 1, no. 12, pp. 1322–1324, 1995. View at: Publisher Site | Google Scholar
  141. M. J. Yaszemski, R. G. Payne, W. C. Hayes, R. Langer, and A. G. Mikos, “Evolution of bone transplantation: molecular, cellular and tissue strategies to engineer human bone,” Biomaterials, vol. 17, no. 2, pp. 175–185, 1996. View at: Publisher Site | Google Scholar
  142. E. M. Younger and M. W. Chapman, “Morbidity at bone graft donor sites,” Journal of Orthopaedic Trauma, vol. 3, no. 3, pp. 192–195, 1989. View at: Publisher Site | Google Scholar
  143. S. Gronthos and P. J. Simmons, “The biology and application of human bone marrow stromal cell precursors,” Journal of Hematotherapy and Stem Cell Research, vol. 5, no. 1, pp. 15–23, 1996. View at: Google Scholar
  144. N. Jaiswal, S. E. Haynesworth, A. I. Caplan, and S. P. Bruder, “Osteogenic differentiation of purified, culture-expanded human mesenchymal stem cells in vitro,” Journal of Cellular Biochemistry, vol. 64, no. 2, pp. 295–312, 1997. View at: Google Scholar
  145. J. E. Aubin, “Osteoprogenitor cell frequency in rat bone marrow stromal populations: role for heterotypic cell-cell interactions in osteoblast differentiation,” Journal of Cellular Biochemistry, vol. 72, no. 3, pp. 396–410, 1999. View at: Google Scholar
  146. J. Rauh, F. Milan, K. Günther, and M. Stiehler, “Bioreactor systems for bone tissue engineering,” Tissue Engineering B: Reviews, vol. 17, no. 4, pp. 263–280, 2011. View at: Publisher Site | Google Scholar
  147. S. Kern, H. Eichler, J. Stoeve, H. Klüter, and K. Bieback, “Comparative analysis of mesenchymal stem cells from bone marrow, umbilical cord blood, or adipose tissue,” Stem Cells, vol. 24, no. 5, pp. 1294–1301, 2006. View at: Publisher Site | Google Scholar
  148. M. Kassem and B. M. Abdallah, “Human bone-marrow-derived mesenchymal stem cells: biological characteristics and potential role in therapy of degenerative diseases,” Cell and Tissue Research, vol. 331, no. 1, pp. 157–163, 2008. View at: Publisher Site | Google Scholar
  149. J. E. Dennis, S. E. Haynesworth, R. G. Young, and A. I. Caplan, “Osteogenesis in marrow-derived mesenchymal cell porous ceramic composites transplanted subcutaneously: effect of fibronectin and laminin on cell retention and rate of osteogenic expression,” Cell Transplantation, vol. 1, no. 1, pp. 23–32, 1992. View at: Google Scholar
  150. B. D. Boyan, T. W. Hummert, D. D. Dean, and Z. Schwartz, “Role of material surfaces in regulating bone and cartilage cell response,” Biomaterials, vol. 17, no. 2, pp. 137–146, 1996. View at: Publisher Site | Google Scholar
  151. A. L. Olivares, È. Marsal, J. A. Planell, and D. Lacroix, “Finite element study of scaffold architecture design and culture conditions for tissue engineering,” Biomaterials, vol. 30, no. 30, pp. 6142–6149, 2009. View at: Publisher Site | Google Scholar
  152. S. B. Vangordon, R. S. Voronov, T. B. Blue, R. L. Shambaugh, D. V. Papavassiliou, and V. I. Sikavitsas, “Effects of scaffold architecture on preosteoblastic cultures under continuous fluid shear,” Industrial & Engineering Chemistry Research, vol. 50, no. 2, pp. 620–629, 2011. View at: Publisher Site | Google Scholar
  153. E. Canalis, A. N. Economides, and E. Gazzerro, “Bone morphogenetic proteins, their antagonists, and the skeleton,” Endocrine Reviews, vol. 24, no. 2, pp. 218–235, 2003. View at: Publisher Site | Google Scholar
  154. B. L. Eppley, J. E. Woodell, and J. Higgins, “Platelet quantification and growth factor analysis from platelet-rich plasma: implications for wound healing,” Plastic and Reconstructive Surgery, vol. 114, no. 6, pp. 1502–1508, 2004. View at: Publisher Site | Google Scholar
  155. E. A. Botchwey, S. R. Pollack, E. M. Levine, and C. T. Laurencin, “Bone tissue engineering in a rotating bioreactor using a microcarrier matrix system,” Journal of Biomedical Materials Research, vol. 55, no. 2, pp. 242–253, 2001. View at: Publisher Site | Google Scholar
  156. J. Chen, R. Liu, Y. Yang et al., “The simulated microgravity enhances the differentiation of mesenchymal stem cells into neurons,” Neuroscience Letters, vol. 505, no. 2, pp. 171–175, 2011. View at: Publisher Site | Google Scholar
  157. D. N. Karunaratne, P. S. Silverstein, V. Vasadani et al., “Cell culture models for drug transport studies,” in Drug Delivery: Principles and Applications, B. Wang, T. Siahaan, and R. Soltero, Eds., pp. 103–124, John Wiley & Sons, Hoboken, NJ, USA, 2005. View at: Google Scholar
  158. M. Gassmann, J. Fandrey, S. Bichet et al., “Oxygen supply and oxygen-dependent gene expression in differentiating embryonic stem cells,” Proceedings of the National Academy of Sciences of the United States of America, vol. 93, no. 7, pp. 2867–2872, 1996. View at: Publisher Site | Google Scholar
  159. R. M. Sutherland, “Cell and environment interactions in tumor microregions: the multicell spheroid model,” Science, vol. 240, no. 4849, pp. 177–184, 1988. View at: Google Scholar
  160. L. A. Kunz-Schughart, J. P. Freyer, F. Hofstaedter, and R. Ebner, “The use of 3-D cultures for high-throughput screening: the multicellular spheroid model,” Journal of Biomolecular Screening, vol. 9, no. 4, pp. 273–285, 2004. View at: Publisher Site | Google Scholar
  161. F. Hirschhaeuser, H. Menne, C. Dittfeld, J. West, W. Mueller-Klieser, and L. A. Kunz-Schughart, “Multicellular tumor spheroids: an underestimated tool is catching up again,” Journal of Biotechnology, vol. 148, no. 1, pp. 3–15, 2010. View at: Publisher Site | Google Scholar
  162. D. D. Fang, Y. J. Kim, C. N. Lee et al., “Expansion of CD133+ colon cancer cultures retaining stem cell properties to enable cancer stem cell target discovery,” British Journal of Cancer, vol. 102, no. 8, pp. 1265–1275, 2010. View at: Publisher Site | Google Scholar
  163. S. M. Ong, C. Zhang, Y. Toh et al., “A gel-free 3D microfluidic cell culture system,” Biomaterials, vol. 29, no. 22, pp. 3237–3244, 2008. View at: Publisher Site | Google Scholar
  164. Y.-S. Torisawa, B.-H. Chueh, D. Huh et al., “Efficient formation of uniform-sized embryoid bodies using a compartmentalized microchannel device,” Lab on a Chip, vol. 7, no. 6, pp. 770–776, 2007. View at: Publisher Site | Google Scholar
  165. L. Y. Wu, D. Di Carlo, and L. P. Lee, “Microfluidic self-assembly of tumor spheroids for anticancer drug discovery,” Biomedical Microdevices, vol. 10, no. 2, pp. 197–202, 2008. View at: Publisher Site | Google Scholar
  166. Y. S. Torisawa, A. Takagi, Y. Nashimoto, T. Yasukawa, H. Shiku, and T. Matsue, “A multicellular spheroid array to realize spheroid formation, culture, and viability assay on a chip,” Biomaterials, vol. 28, no. 3, pp. 559–566, 2007. View at: Publisher Site | Google Scholar
  167. J. M. Kelm and M. Fussenegger, “Microscale tissue engineering using gravity-enforced cell assembly,” Trends in Biotechnology, vol. 22, no. 4, pp. 195–202, 2004. View at: Publisher Site | Google Scholar
  168. J. Friedrich, R. Ebner, and L. A. Kunz-Schughart, “Experimental anti-tumor therapy in 3-D: spheroids—old hat or new challenge?” International Journal of Radiation Biology, vol. 83, no. 11-12, pp. 849–871, 2007. View at: Publisher Site | Google Scholar
  169. J. M. Kelm and M. Fussenegger, “Scaffold-free cell delivery for use in regenerative medicine,” Advanced Drug Delivery Reviews, vol. 62, no. 7-8, pp. 753–764, 2010. View at: Publisher Site | Google Scholar
  170. V. M. Weaver, O. W. Petersen, F. Wang et al., “Reversion of the malignant phenotype of human breast cells in three-dimensional culture and in vivo by integrin blocking antibodies,” The Journal of Cell Biology, vol. 137, no. 1, pp. 231–245, 1997. View at: Publisher Site | Google Scholar
  171. M. Hughes-Fulford, “Function of the cytoskeleton in gravisensing during spaceflight,” Advances in Space Research, vol. 32, no. 8, pp. 1585–1593, 2003. View at: Publisher Site | Google Scholar
  172. Y. Naito, T. Shinoka, D. Duncan et al., “Vascular tissue engineering: towards the next generation vascular grafts,” Advanced Drug Delivery Reviews, vol. 63, no. 4-5, pp. 312–323, 2011. View at: Publisher Site | Google Scholar
  173. C. Kang, L. Zou, M. Yuan et al., “Impact of simulated microgravity on microvascular endothelial cell apoptosis,” European Journal of Applied Physiology, vol. 111, no. 9, pp. 2131–2138, 2011. View at: Publisher Site | Google Scholar
  174. S. M. Grenon, M. Jeanne, J. Aguado-Zuniga, M. S. Conte, and M. Hughes-Fulford, “Effects of gravitational mechanical unloading in endothelial cells: association between caveolins, inflammation and adhesion molecules,” Scientific Reports, vol. 3, article 1494, 2013. View at: Publisher Site | Google Scholar

Copyright © 2014 Claudia Ulbrich et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

More related articles

 PDF Download Citation Citation
 Download other formatsMore
 Order printed copiesOrder

Related articles

We are committed to sharing findings related to COVID-19 as quickly as possible. We will be providing unlimited waivers of publication charges for accepted research articles as well as case reports and case series related to COVID-19. Review articles are excluded from this waiver policy. Sign up here as a reviewer to help fast-track new submissions.