ALLS: Artificial Life-Like Systems

Design of Artificial Cells from Functional Modules

Synthetic Biology is an exciting and rapidly evolving field focused on engineering cells and biological systems to perform predictable and tailored functions—whether modified, enhanced, or entirely novel. The approach involves breaking down the inherent complexity of biological systems into smaller subsystems and functional modules, which are then redesigned and reassembled into larger, integrated units. This methodology is often compared to the design of biotechnological or chemical plants (Rollié et al., 2012). In this context, the PSE Group contributed its expertise in assembling and controlling chemical, energy, and production processes to the MaxSynBio initiative. This research consortium brought together eight Max Planck Institutes, along with research groups from Friedrich-Alexander University Erlangen-Nuremberg and the University of Bordeaux. It was jointly funded by the Max Planck Society and the Federal Ministry of Education and Research in Germany. Building on the achievements of MaxSynBio, the PSE Group’s work in synthetic cell research has progressed into the Artificial Life-like Systems (ALLS) Team, continuing the pursuit of engineered life-like systems with controllable and modular functionalities.

Synthetic Cell Engineering in the ALLS Team

The primary scientific goal of the ALLS Team is to develop a true bottom-up engineering approach (Fig. 1) to mimic the fundamental structural and functional principles of living cells. This approach not only advances our understanding of biological systems but also enables the development of novel synthetic platforms. While the near future of synthetic biology is expected to continue relying largely on top-down strategies (i.e., constructing minimal systems from existing organisms), bottom-up approaches—starting from first principles—offer the potential to refine and enhance these systems using synthetic or alternative biological components, paving the way for a new generation of biotechnological processes.

Fig. 1: Workflow for the bottom-up assembly of an artificial mitochondrion. Left: characterization of functional parts through performance standards such as activity, stability, etc.; Middle: integration of functional parts into functional modules for metabolism, energy and transport through assembly standards; Right: integration of modules into hypothetical artificial mitochondrion system through assembly standards and computer-aided optimization. — **Fig. 1:** Workflow for the bottom-up assembly of an artificial mitochondrion. Left: characterization of functional parts through performance standards such as activity, stability, etc.; Middle: integration of functional parts into functional modules for metabolism, energy and transport through assembly standards; Right: integration of modules into hypothetical artificial mitochondrion system through assembly standards and computer-aided optimization.

**Fig. 1:** Workflow for the bottom-up assembly of an artificial mitochondrion. Left: characterization of functional parts through performance standards such as activity, stability, etc.; Middle: integration of functional parts into functional modules for metabolism, energy and transport through assembly standards; Right: integration of modules into hypothetical artificial mitochondrion system through assembly standards and computer-aided optimization.

Researchers in the PSE group have defined the minimal functional modules necessary for creating artificial life-like systems, with a strong focus on inter-module compatibility (Fig. 2). These core functionalities include:

Fig. 2: Schematic illustration of the artificial cell architecture as designed by ALLS Team, highlighting the key structural components and functional elements. — **Fig. 2:** Schematic illustration of the artificial cell architecture as designed by ALLS Team, highlighting the key structural components and functional elements.

**Fig. 2:** Schematic illustration of the artificial cell architecture as designed by ALLS Team, highlighting the key structural components and functional elements.

Motility
Sensing/Signaling
Metabolism
Transport
Energy generation

For instance, motility enables artificial cells to move in response to environmental stimuli, mimicking key aspects of living organisms. This is tightly integrated with the sensing/signaling module. The energy required for activating motility is supplied in the form of ATP, produced by a metabolic cascade and an ATP-generating module. Enabling technologies such as compartmentalization, membrane transport, cell-free protein expression, and membrane fusion are under continuous development to support these modules.

Importantly, our research is not limited to mimicking natural functions. By integrating natural and synthetic materials, we are uncovering novel and unexpected behaviors in hybrid systems. One key advancement has been the replacement of natural membrane components (e.g., lipids) with synthetic amphiphilic copolymers, which offer broad chemical versatility and tunability for applications in technology, biology, and medicine.

To form vesicles from these synthetic amphiphiles, we are developing and optimizing several fabrication techniques and functionalization protocols, including:

Gentle film rehydration and extrusion
Electroformation (and fusion-assisted electroformation)
Droplet phase transfer
Microfluidics

For membrane protein functionalization of lipid, polymer, or hybrid membranes, we commonly use:

Detergent-mediated reconstitution
Droplet phase transfer
Membrane fusion

Asymmetric Membranes and Natural-Synthetic Compatibility

Fig. 3: Reconstituting minimal phosphorylation machinery in polymer and hybrid membranes. Above: homogenous lipid/polymer hybrid membrane in microcontainers; Below: respiratory-driven ATP synthesis in lipid, hybrid and polymer nanocontainers. — **Fig. 3:** Reconstituting minimal phosphorylation machinery in polymer and hybrid membranes. Above: homogenous lipid/polymer hybrid membrane in microcontainers; Below: respiratory-driven ATP synthesis in lipid, hybrid and polymer nanocontainers.

**Fig. 3:** Reconstituting minimal phosphorylation machinery in polymer and hybrid membranes. Above: homogenous lipid/polymer hybrid membrane in microcontainers; Below: respiratory-driven ATP synthesis in lipid, hybrid and polymer nanocontainers.

Fig. 4: Schematic illustration of different types of polymer membrane architectures: AB diblock copolymers with transmembrane confirmation, ABA triblock copolymers where architecture is a mixture of transmembrane and hairpins, which have both their hydrophilic blocks displayed at the same membrane surface, ABC triblock copolymers with transmembrane confirmation, graft copolymer with hydrophobic backbone in hairpin conformation and hydrophilic side chains facing outwards, Janus dendrimers forming membrane of dendrimersomes. — **Fig. 4:** Schematic illustration of different types of polymer membrane architectures: AB diblock copolymers with transmembrane confirmation, ABA triblock copolymers where architecture is a mixture of transmembrane and hairpins, which have both their hydrophilic blocks displayed at the same membrane surface, ABC triblock copolymers with transmembrane confirmation, graft copolymer with hydrophobic backbone in hairpin conformation and hydrophilic side chains facing outwards, Janus dendrimers forming membrane of dendrimersomes.

**Fig. 4:** Schematic illustration of different types of polymer membrane architectures: AB diblock copolymers with transmembrane confirmation, ABA triblock copolymers where architecture is a mixture of transmembrane and hairpins, which have both their hydrophilic blocks displayed at the same membrane surface, ABC triblock copolymers with transmembrane confirmation, graft copolymer with hydrophobic backbone in hairpin conformation and hydrophilic side chains facing outwards, Janus dendrimers forming membrane of dendrimersomes.

Fig. 5: Protein-functionalized microcompartments. (A) Bending rigidity for protein-free (w/o bo3; white area) and protein-functionalized (w/ bo3; gray area) liposomes, hybrids, and polymersomes. (B) bo3-polymer-GUVs with encapsulated pH-sensitive dye (green) trapped in microfluidic chip. Scale bar, 50 μm. (C) Successful insertion and homogenous distribution of bo3 oxidase-ATTO 514 (green) in bo3-polymer-GUVs. Polymer labeled with rhodamine (PDMS-g-PEO-Rho, red) was used to visualize the memrbane. Scale bar, 10 μm. — **Fig. 5:** Protein-functionalized microcompartments. (A) Bending rigidity for protein-free (w/o *bo₃*; white area) and protein-functionalized (w/ *bo₃*; gray area) liposomes, hybrids, and polymersomes. (B) *bo₃*-polymer-GUVs with encapsulated pH-sensitive dye (green) trapped in microfluidic chip. Scale bar, 50 μm. (C) Successful insertion and homogenous distribution of *bo₃* oxidase-ATTO 514 (green) in *bo₃*-polymer-GUVs. Polymer labeled with rhodamine (PDMS-g-PEO-Rho, red) was used to visualize the memrbane. Scale bar, 10 μm.

**Fig. 5:** Protein-functionalized microcompartments. (A) Bending rigidity for protein-free (w/o *bo₃*; white area) and protein-functionalized (w/ *bo₃*; gray area) liposomes, hybrids, and polymersomes. (B) *bo₃*-polymer-GUVs with encapsulated pH-sensitive dye (green) trapped in microfluidic chip. Scale bar, 50 μm. (C) Successful insertion and homogenous distribution of *bo₃* oxidase-ATTO 514 (green) in *bo₃*-polymer-GUVs. Polymer labeled with rhodamine (PDMS-g-PEO-Rho, red) was used to visualize the memrbane. Scale bar, 10 μm.

Biological membranes are highly asymmetric, a property essential for functions such as signal transduction. While achieving and maintaining lipid asymmetry is challenging, we explore asymmetric polymer membranes for specific applications. Giant unilamellar vesicles (GUVs) with asymmetric membranes are assembled using either a modified droplet phase transfer method or double-emulsion microfluidics, where each leaflet is formed from a different polymer.

We are particularly interested in the integration of natural components from diverse organisms with synthetic membranes—specifically, the functional behavior of membrane proteins in polymer-based environments (Fig. 3; Otrin et al., 2017; Marušič et al., 2020). We are also exploring various polymer membrane architectures to assess their biocompatibility with natural elements (Fig. 4).

To this end, we have developed tools to study membrane protein activity at the single-vesicle level (Fig. 5), assessing key properties such as: membrane order/disorder, lateral diffusion, thermal stability, and permeability.

Membrane Dynamics and Energy Modules

Understanding membrane dynamics is essential for replicating cellular functions like endocytosis, exocytosis, and division. Our lab explores fusion and fission mechanisms using various strategies, including:

Mechanical stress
Salt-induced fusion
Electrostatic interactions (via charged lipids/polymers)
Fusogenic peptides (e.g., SNARE proteins)

Fig. 6. A schematic representation of light-driven (A) and oxidative phosphorylation-based chemically-driven (B) ATP regeneration in compartments. Proton motive force, driving the synthesis of ATP by the ATP synthase, is, in light-driven systems, generated by either oxidation of water by photosystem II (PSII) coupled to the reduction of electron mediator (MED) (A1) or by proton translocation via light-driven proton pump, such as bacteriorhodopsin (bRH) (A3). Alternatively, transmembrane proton gradient can be established by so-called photoacid (PAOH) or photobase(PBOH) generators, triggered by light. In comparison, chemically-driven systems rely solely on proton translocation coupled with various redox reactions between electron donors (ED) and electron acceptors (EA). While the examples B1 and B2 are essentially very similar, we showcase the possibility of sodium-driven systems and well as simultaneous NAD+ regeneration. — **Fig. 6.** A schematic representation of light-driven (A) and oxidative phosphorylation-based chemically-driven (B) ATP regeneration in compartments. Proton motive force, driving the synthesis of ATP by the ATP synthase, is, in light-driven systems, generated by either oxidation of water by photosystem II (PSII) coupled to the reduction of electron mediator (MED) (A1) or by proton translocation via light-driven proton pump, such as bacteriorhodopsin (bRH) (A3). Alternatively, transmembrane proton gradient can be established by so-called photoacid (PAOH) or photobase(PBOH) generators, triggered by light. In comparison, chemically-driven systems rely solely on proton translocation coupled with various redox reactions between electron donors (ED) and electron acceptors (EA). While the examples B1 and B2 are essentially very similar, we showcase the possibility of sodium-driven systems and well as simultaneous NAD⁺ regeneration.

**Fig. 6.** A schematic representation of light-driven (A) and oxidative phosphorylation-based chemically-driven (B) ATP regeneration in compartments. Proton motive force, driving the synthesis of ATP by the ATP synthase, is, in light-driven systems, generated by either oxidation of water by photosystem II (PSII) coupled to the reduction of electron mediator (MED) (A1) or by proton translocation via light-driven proton pump, such as bacteriorhodopsin (bRH) (A3). Alternatively, transmembrane proton gradient can be established by so-called photoacid (PAOH) or photobase(PBOH) generators, triggered by light. In comparison, chemically-driven systems rely solely on proton translocation coupled with various redox reactions between electron donors (ED) and electron acceptors (EA). While the examples B1 and B2 are essentially very similar, we showcase the possibility of sodium-driven systems and well as simultaneous NAD⁺ regeneration.

Active processes in living systems require continuous energy and material supply, tightly linked to cellular metabolism. Following the bottom-up approach, we have assembled and characterized fundamental components for energy conversion and transduction, such as light- and chemically-driven proton pumps and ATP synthase (Fig. 6). We are continuously expanding our library of purified membrane proteins, including oxidases, porins, and fusogenic peptides.

Our expertise includes the assembly and characterization of proteins involved in oxidative phosphorylation and photophosphorylation, including assessments of protein orientation and enzymatic activity. We have also demonstrated cofactor regeneration via encapsulated hydrophobic redox mediators, mimicking the functions of NADH dehydrogenase and cytochrome c reductase (Fig. 7; Wang et al., 2018).

Fig. 7: NAD regeneration module. (A) Complexes I and III – natural functional parts for NADH oxidation and transmembrane electron transfer; (B) a synthetic analogue inserted into phospholipid bilayer. — **Fig. 7:** NAD regeneration module. (A) Complexes I and III – natural functional parts for NADH oxidation and transmembrane electron transfer; (B) a synthetic analogue inserted into phospholipid bilayer.

**Fig. 7:** NAD regeneration module. (A) Complexes I and III – natural functional parts for NADH oxidation and transmembrane electron transfer; (B) a synthetic analogue inserted into phospholipid bilayer.

Fig. 8: Proposed fusion intermediates of SNARE-mediated fusion in hybrids. Long contact is established between the lipid-rich domain (bilayer) of one fusing vesicle, and the polymer-rich domain of another (I). The initial contact between the fusing membranes occurs at the lateral edge of the protruding bilayer and it further develops into crosswise membrane mixing (the process of “hybrid zippering”, II). Transient fusion diaphragm, exhibiting lipid-rich bilayer stabilized in polymer bilayer is observed (III) before stable lipid bilayers are formed in the newly mixed hybrid diaphragm (IV). The fusion pore was formed in the more central region of the juncture (V). No inclusions or deformations can be observed in fused vesicles (VI). Scale bars, 30 nm. — **Fig. 8:** Proposed fusion intermediates of SNARE-mediated fusion in hybrids. Long contact is established between the lipid-rich domain (bilayer) of one fusing vesicle, and the polymer-rich domain of another (I). The initial contact between the fusing membranes occurs at the lateral edge of the protruding bilayer and it further develops into crosswise membrane mixing (the process of “hybrid zippering”, II). Transient fusion diaphragm, exhibiting lipid-rich bilayer stabilized in polymer bilayer is observed (III) before stable lipid bilayers are formed in the newly mixed hybrid diaphragm (IV). The fusion pore was formed in the more central region of the juncture (V). No inclusions or deformations can be observed in fused vesicles (VI). Scale bars, 30 nm.

**Fig. 8:** Proposed fusion intermediates of SNARE-mediated fusion in hybrids. Long contact is established between the lipid-rich domain (bilayer) of one fusing vesicle, and the polymer-rich domain of another (I). The initial contact between the fusing membranes occurs at the lateral edge of the protruding bilayer and it further develops into crosswise membrane mixing (the process of “hybrid zippering”, II). Transient fusion diaphragm, exhibiting lipid-rich bilayer stabilized in polymer bilayer is observed (III) before stable lipid bilayers are formed in the newly mixed hybrid diaphragm (IV). The fusion pore was formed in the more central region of the juncture (V). No inclusions or deformations can be observed in fused vesicles (VI). Scale bars, 30 nm.

Optimizing membrane protein insertion requires extensive screening of reconstitution conditions, which often vary significantly across different enzymes. To address this, we explore simultaneous integration of multiple enzymes via membrane fusion, utilizing SNARE proteins. Fusion intermediates between polymer and hybrid membranes have also been characterized (Fig. 8; Otrin et al., 2021).

In parallel, we explore metabolic pathways and their integration with energy modules. We also utilize the E. coli respiratory chain as a natural reference system, combining it with minimal synthetic metabolism (Beneyton et al., 2018). Our energy and metabolic modules are further described using mathematical modeling to enable system optimization and provide a quantitative framework for integration.

Finally, to enable targeted localization of signaling and motility modules, we investigate lateral phase separation (Fig. 9; Otrin et al., 2024) in membranes composed of multiple components (e.g., lipids and polymers or polymer blends). We achieve control over microscopically heterogeneous or homogeneous membranes through:

Membrane composition ratios
Differences in hydrophobic thickness
Temperature modulation
Electrostatic interactions
Membrane protein integration

Fig. 9: Membrane proteins partitioning in polymer/lipid hybrid GUVs. Upper panel: homogenous distribution of proteins; lower panel: proteins partitioning in lipid domains. Proton pump bo3 oxidase is labeled with ATTO 514 (green), ATTO 520 (yellow) or ATTO 425 (cyan) and F1FO-ATPase with ATTO 620 (magenta). Scale bar, 5 μm. — **Fig. 9**: Membrane proteins partitioning in polymer/lipid hybrid GUVs. Upper panel: homogenous distribution of proteins; lower panel: proteins partitioning in lipid domains. Proton pump *bo₃* oxidase is labeled with ATTO 514 (green), ATTO 520 (yellow) or ATTO 425 (cyan) and F₁F_O-ATPase with ATTO 620 (magenta). Scale bar, 5 μm.

**Fig. 9**: Membrane proteins partitioning in polymer/lipid hybrid GUVs. Upper panel: homogenous distribution of proteins; lower panel: proteins partitioning in lipid domains. Proton pump *bo₃* oxidase is labeled with ATTO 514 (green), ATTO 520 (yellow) or ATTO 425 (cyan) and F₁F_O-ATPase with ATTO 620 (magenta). Scale bar, 5 μm.

Recent Publications

Otrin N., Otrin L., Bednarz C., Träger T. K., Hamdi F., Kastritis P. L., Ivanov I., and Sundmacher K. (2024). Protein-rich rafts in hybrid polymer/lipid giant unilamellar vesicles. Biomacromolecules 25(2): 778-791

Marušič N., Otrin L., Rauchhaus J., Zhao Z., Kyrilis F.L., Hamdi F., Kastritis P.L., Dimova R., Ivanov I., and Sundmacher K. (2022). Increased efficiency of charge-mediated fusion in polymer/lipid hybrid membranes. Proceedings of National Academy of Sciences 119(20): e2122468119

Marušič N., Zhao Z., Otrin L., Dimova R., Ivanov I., and Sundmacher K. (2022). Fusion-induced Growth of Biomimetic Polymersomes: Behavior of poly(dimethylsiloxane)-poly(ethylene oxide) Vesicles in Saline Solutions Under High Agitation. Macromolecular Rapid Communications 43(5): e2100712

Staufer O., De Lora J. A., Bailoni E., Bazrasfhan A., Benk A. S., Jahnke K., Manzer Z. A., Otrin L., Pérez T. D., Sharon J., Steinkühler J., Adamala K. P., Jacobson B., Dogterom M., Göpfrich K., Stefanovic D., Atlas S. R., Grunze M., Lakin M. R., Shreve A. P., Spatz J. S., and López G. P. (2021). Science Forum: Building a community to engineer synthetic cells and organelles from the bottom-up. eLife 10: e73556

Otrin L., Witkowska A., Marušič N., Zhao Z., B. Lira R., Kyrilis F., Hamdi F., Ivanov I., Lipowsky R., L. Kastritis P., Dimova R., Sundmacher K., Jahn R., and Vidaković-Koch T. (2021). En route to dynamic life processes by SNARE-mediated fusion of polymer and hybrid membranes. Nature Communications 12: 4972

Shetty C. S., Yandrapalli N., Pinkwart K., Krafft D., Vidakovic-Koch T., Ivanov I., and Robinson T. (2021). Directed Signaling Cascades in Monodisperse Artificial Eukaryotic Cells. ACS Nano 15: 15656−15666

Balasbas III S., Ivanov I. and Sundmacher K. (2021). Metabolic Network Reconstruction from Time-Series of Concentration Data: Evaluation of the Numerical Matrices Methods. Computer Aided Chemical Engineering 50: 1991-1996

Ivanov I., López Castellanos S., Balasbas III S., Otrin L., Marušič N., Vidaković-Koch T., and Sundmacher K. (2021). Bottom-Up Synthesis of Artificial Cells: Recent Highlights and Future Challenges. Annual Review of Chemical and Biomolecular Engineering 12: 287-308

Wang M., Weber A., Hartig R., Zheng Y., Krafft D., Vidaković-Koch T., Zuschratter W., Ivanov I., Sundmacher K. (2021). Scale up of transmembrane NADH oxidation in synthetic giant vesicles. Bioconjugate Chemistry 32 (5): 897–903.

Ahmad R., Kleineberg C., Nasirimarekani V., Su Y.-J., Pozveh S. G., Bae A., Sundmacher K., Bodenschatz E., Guido I., Vidaković-Koch T., and Gholami A. (2021). Light-Powered Reactivation of Flagella and Contraction of Microtubule Networks: Toward Building an Artificial Cell. ACS Synthetic Biology 10 (6): 1490–1504

Wohlfromm F., Richter M., Otrin L., Seyrek K., Vidaković-Koch T., Kuligina E., Richter V., Koval O., and Lavrik I. N. (2021). Interplay Between Mitophagy and Apoptosis Defines a Cell Fate Upon Co-treatment of Breast Cancer Cells With a Recombinant Fragment of Human κ-Casein and Tumor Necrosis Factor-Related Apoptosis-Inducing Ligand. Frontiers in Cell and Developmental Biology 8: 617762

Xu D., Kleineberg C., Vidaković-Koch T., Wegner S. V. (2020). Multistimuli sensing adhesion unit for the self-positioning of minimal synthetic cells. Small 16 (35): 2002440

Marušič N., Otrin O., Zhao Z., Lira R.B., Kyrilis F.L., Hamdi F., Kastritis P.L., Vidaković-Koch T., Ivanov I., Sundmacher K., and Dimova R. (2020). Constructing artificial respiratory chain in polymer compartments: Insights into the interplay between bo₃ oxidase and the membrane. Proceedings of National Academy of Sciences 117 (26): 15006-15017

Kleineberg C., Wölfer C., Abbasnia A., Pischel D., Bednarz C., Ivanov I., Heitkamp T., Börsch M., Sundmacher K., Vidaković-Koch T. (2020). Light-driven ATP regeneration in diblock/grafted hybrid vesicles. ChemBioChem 21 (15): 2149-2160

Otrin L., Kleineberg C., de Silva L.C., Landfester K., Ivanov I., Wang M, Bednarz C., Sundmacher K., Vidaković-Koch T. (2019). Artifical organelles for energy regeneration. Advanced Biosystems 3 (6): 1800323

Ivanov I., Lira R.B., Tang T.-Y.D., Franzmann T., Klosin A., de Silva L.C., Hyman A., Landfester K., Lipowsky R., Sundmacher K., and Dimova R. (2019). Directed growth of biomimetic microcompartments. Advanced Biosystems 3: 1800314

Krafft D., López Castellanos S., Lira R.B., Dimova R., Ivanov I., Sundmacher K. (2019). Compartments for synthetic cells: Osmotically assisted separtion of oil from double emulsions in microfluidic chip. ChemBioChem 20: 2604-2608

Beneyton T., Krafft D., Bednarz C., Kleineberg C.,Woelfer C., Ivanov I., Vidaković-Koch T., Sundmacher K., and Baret J. C. (2018). Out-of-equilibrium microcompartments for the bottom-up integration of metabolic functions. Nature Communications 9: 2391

Wang M., Wölfer C., Otrin L., Ivanov I., Vidaković-Koch T., and Sundmacher K. (2018). Transmembrane NADH Oxidation with Tetracyanoquinodimethane. Langmuir 34 (19): 5435-5443

Weiss M., Frohnmayer J. P., Benk L. T., Haller B., Janiesch J. W., Heitkamp T., Börsch M., Lira R. B., Dimova R., Lipowsky R., Bodenschatz E., Baret J. C., Vidakovic-Koch T., Sundmacher K., Platzman I., Spatz J. P. (2018). Sequential bottom-up assembly of mechanically stabilized synthetic cells by microfluidics. Nature Materials 17(1): 89-96

Schwille P., Spatz J., Landfester K., Bodenschatz E., Herminghaus S., Sourjik V., Erb T. J., Bastiaens P., Lipowsky R., Hyman A., Dabrock P., Baret J. C., Vidaković-Koch T., Bieling P., Dimova R., Mutschler H., Robinson T., Tang T. D., Wegner S., and Sundmacher K. (2018). MaxSynBio: Avenues Towards Creating Cells from the Bottom Up. Angewandte Chemie International Edition 57 (41):13382-13392

Otrin L., Marušič N., Bednarz C., Vidaković-Koch T., Lieberwirth I., Landfester K., and Sundmacher K. (2017). Toward Artificial Mitochondrion: Mimicking Oxidative Phosphorylation in Polymer and Hybrid Membranes. Nano Letters 17 (11): 6816-6821

Rollié S., Mangold M. and Sundmacher K. (2012). Designing biological systems: systems engineering meets synthetic biology. Chemical Engineering Science, 69 (1): 1-29

Collaborations

Natural Building Blocks

Prof. Volker Zickermann, Medical Faculty of Goethe University, Frankfurt, Germany

Prof. Marc Nowaczyk, Ruhr University Bochum, Bochum, Germany

Prof. Sonja-Verena Albers, University of Freiburg, Freiburg, Germany

Synthetic Building Blocks

Prof. Sebastién Lecommandoux, Université de Bordeaux, Bordeaux, France

Prof. César Rodriguez-Emmenegger, Institute for Bioengineering of Catalonia, Barcelona, Spain

Biophysical and Structural Analysis of Vesicles

Prof. Panagiotis L. Kastritis, Martin Luther University Halle-Wittenberg, Halle, Germany

Prof. Katharina Landfester, Max Planck Institute for Polymer Research, Mainz, Germany

Dr. Rumiana Dimova, Prof. Reinhard Lipowsky, Max Planck Institute of Colloids and Interfaces, Potsdam, Germany