Despite the undeniable importance of studying natural substrate microbial communities, our understanding of their organization and functioning remains extremely insufficient [1, 2]. The lack of information on soil microbial communities is a consequence of their extreme complexity and genetic diversity [3]. Microbial communities are living systems that are influenced by external factors, which leads to adequate changes in the community, both qualitative and quantitative. Therefore, the study of the functioning of such a dynamic system as substrate microbial communities, the influence of various external factors on them, including anthropogenic ones, is of great importance [4]. Monitoring microorganisms in their natural habitat is necessary for a better understanding of microbial ecosystems and their survival strategies, which is important, among other things, for assessing the risk of introducing genetically modified microorganisms into natural communities [5]. For the study of microbial communities, classical microbiological methods have been used for a long time. Namely: methods of making substrate (soil) washes, sowing these washes on nutrient media, isolating the resulting colonies by morphotypes into pure cultures with their further study. However, it soon became known that not all microorganisms are capable of growing on artificial nutrient media. In this regard, I would like to especially note the emphasis made by Akkermans [5]. on the fact that the concept of "unculturable" in relation to microorganisms is not correct as long as their inability to culture on artificial nutrient media is actually proven. In general, the author considers it appropriate to use the term "uncultured", meaning that this microorganism cannot be cultivated at the moment due to the lack of technology for its cultivation, but hypothetically it can be obtained under artificial conditions. Really, a number of studies show that some uncultured microorganisms become cultureable under specific experimental conditions Kaeberiein Joseph. Thus, scientists have come to understand that microorganisms that can be isolated from the microbial community on separate nutrient media are only an insignificant part of the community as in total Kaeberlein Torre. Microbial cenoses are very heterogeneous and consist of cells with a wide variety of metabolic abilities. In order to describe such biodiversity, realistic, reliable and accurate methods are required that allow both the detection and identification of microbial groups. Such methods have appeared. These methods of molecular analysis that study microcenoses at the molecular level allow working with uncultured forms [3]. Such methods have really opened up fundamentally new possibilities in the study of cenoses. However, like the previously used classical cultivation methods, they were destructive. That is, the application of such approaches includes the complete destruction of the microcenosis and the extraction of its components (cells in the case of culture methods, nucleic acids in the case of genetic molecular analysis, lipids and enzymes in the case of biochemical tests). Nevertheless, in the literature, the study and identification of individual structural components is called “studying the structure of microbial communities” [3]. In addition, this is only because today there are no approaches to working with microbial communities in a natural unique structure without disrupting it. Thus, the mechanism of functioning of substrate (both natural and artificial) cenoses is today an unsolved, albeit quite urgent, problem. Today, the substrates in which plants grow can be divided into two large groups (unequal in distribution) - natural and artificial. This division is rather arbitrary, since artificial substrates are mainly made from natural ones. Nevertheless, in the generally accepted understanding, such a division is justified, because the "line of separation" runs between "what is in nature" and "what is combined with the technologies of preparation and then the use of artificial substrates". With the preparation of the substrate (either natural or artificial), the cultivation of plants begins Methods . From that moment on, the problem of microorganisms arises, namely the stable functioning of substrate microcenoses. In the overwhelming majority of cases, the process of formation of substrate microcenoses starts to flow and proceeds out of control, according to its own laws. For natural substrates, under normal natural conditions, such self-organizing microbial cenoses function successfully (although much attention is paid to their knowledge and attempts to control them) [6]. For artificial substrates that do not have stable formed microcenoses at the time of use, the problem of microcenosis organization (directed or spontaneous) often requires special solutions. Therefore, in the case of the formation of a microcenosis in an artificial substrate, control and monitoring of such a rhizosphere cenosis at the stages of its formation and functioning is simply necessary. However, the ability to monitor microorganisms has a number of fundamental limitations. The first of them is associated with the nature of substrates and their structure (Ming, 1989; McGilloway. With the exception of a purely “water” plant growing technology (in which the roots are completely in a nutrient solution), all others use granular-structured substrates. It applies to both natural and artificial (vermiculite, zeolite) substrates. They have developed surfaces consisting of pores, cracks, and cavities in which microorganisms can (and indeed are) located in an amount and quality that is practically inaccessible to research . Even an extremely “open” substrate such as sand has an uneven, complex surface of  sand grains. As a result, due to the fundamental characteristics of the substrate, it fundamentally does not allow for complete monitoring of microorganisms. Also, the problem arises when studying the very biology of the coexistence of microorganisms with a plant. The term "rhizosphere" carries a very clearly defined semantic load, inextricably linked with plant ecology; Whalley at al. 2005). In nature, plants live in natural substrates that form over a long period of time and acquire a definite structure; [7]. For example, the most common substrate for plants is soil. The soil-forming process has been going on for centuries. The result is a mass with a unique rheology. The roots in the soil are surrounded by tightly adhering grains and amorphous material Watteau . In such a substrate, the microflora is organized in a spatial gradient [8] There are microorganisms that exist in the gradient of root secretions directly on the roots. The concentrated maximum of such a gradient falls on the root surface and drops sharply with distance from the root. The drop in the gradient is caused by the diffusion of root exudates into a loosely continuous substrate, which is the soil, as well as the continuous absorption of available organic matter by the entire soil microflora. A consequence of this spatial organization of the plant in the substrate is the organization of the substrate microflora. Root microorganisms reside directly on the roots, basal are located in the first millimeters from the roots, a little further there is a rhizosphere microflora. In turn, the root microflora is also located along the gradient of concentrations of nutrients and phytoncides. Fundamentally the same plant-substrate organization takes place in other habitats [9]. A qualitatively different spatial organization of microflora takes place in artificial substrates - zeolite, vermiculite, etc. There is no loose-continuous medium in them. Such a spatial organization of the community will inevitably lead to a change in the microbial composition.  As there is no such situation in natural communities, there is no ready-made microbial composition for such artificial substrates either. By itself, it will form indefinitely. It should be formed in advance by a series of passages through the same substrate of the required plant, moreover, such an effect, well known for a number of monocultures, as “soil fatigue” should be avoided. The second limitation is due to the nature, biology of the inhabitants of microcenoses, the overwhelming majority of which cannot grow on artificial nutrient media in laboratories. Currently, despite all the efforts and perfection of research methods, no more than 0.1% which is the limiting value, since a figure of 0.01% is usually given) of all microorganisms determined in substrates by direct methods of calculating the total number can grow by the totality of all those culture media that are used in laboratories [10]. To some extent, this is made up for by molecular diagnostic methods - real-time PCR, fluorescent probes, but only to “a certain extent”. The third limitation is determined by what is attributed to the concept of "ecology". For microcenoses, their ecology is almost never defined in terms of reality Vinogradskiy Perfil'ev and Gabe, 1969; Buckey, Schmidt. Over the past 100 years, efforts have been made to overcome these limitations. These include fouling glasses proposed by S.N. Vinogradsky, plane-parallel capillaries by B.V. [11-14]. When solving a number of issues, these methods have not become widespread, because they could not be combined with the highly informative research technologies that emerged in the second half of the last century (electron microscopy, molecular diagnostics, etc.). In addition, in a purely procedural manner, it was inconvenient to work with hard, brittle material. Some brilliant ideas have not found wide application due to the limited technological level of their implementation. We tried, based on modern capabilities, to create the necessary technology that combines the ability to work with microbial cenosis in its natural architecture using all modern techniques with different levels of resolution. The main goal was to develop a non-destructive technology that allows working with cenosis in its native architecture.

Figure 1: Microbial landscape of the rhizosphere zone of Arabidopsis. Luminescence microscopy.

Figure 2: Septic algae of the rhizosphere zone of Arabidopsis. Early stage of formation. Luminescence microscopy.

Figure 3: Microbial-fungal association of the rhizosphere zone of Arabidopsis. Late stage of formation. Scanning electron microscopy.

Figure 4: Microbial-fungal association of the rhizosphere zone of Arabidopsis. Late stage of formation. Luminescencemicroscopy.

Soil was used to form a stable microbial cenosis in an artificial substrate. The soil was taken from the Goloseevsky forest (Kiev, Ukraine). The substrate was placed in a container with soil (10 liters). The residence time of the substrate in the soil is three weeks at a temperature of +23 +25 º? and room humidity. At the same time, the soil was moistened once every three days. As a result, organic substances adhered to the surface of the fibers after three weeks and microbial associations were formed. The plant chosen for the work was Arabidopsis thaliana, cv Columbia. A mini greenhouse was constructed for growing plants. It was a rack with micro-cultivators of plants. Watering with tap water was carried out every two days. The optimum level of atmospheric humidity inside the chamber was 75–80%. To maintain this level, the chambers were covered with transparent plastic lids. Lamps placed over the plants created illumination in the range of 9–12 thousand lux. (at the level of the substrate surface). The system had temperature sensors to control the air temperature both in individual chambers and throughout the system. The temperature regime was within the range of +22 +24 º?. Temperature control was carried out using an air conditioner. The seeds of Arabidopsis thaliana plants were sown to a depth of 1–3 mm. To study the spatial structure of the rhizosphere microbial cenosis at different stages of plant growth, individual fibers were removed from the substrate. Due to the fact that they are placed in parallel, they were easily separated from each other, while maintaining the integrity of the microbial association formed on the surface of the fibers. Each individual fiber was fixed in vapors of 37% formalin for one hour. The fiber was colored with acridine orange dye (Standard Fluka) at a concentration of 0.2% for 5 minutes, washed in tap water. After that, the fiber was placed on a glass slide, covered with a cover glass and examined using fluorescent microscopy. Scanning was carried out over the entire length of the fiber, namely to the entire depth of the substrate. We used an ML-2 fluorescent microscope, LOMO with a light filter with a peak of 380 nm. The fields of view were photographed sequentially, with overlapping. Later the photos were combined into a whole panorama (Adobe Photoshop CS, version 8.0). The same fiber, after studying it by luminescence light microscopy, was analyzed by scanning electron microscopy. For this, the fiber was dried in a desiccator with a ground lid containing 80 silica gel balls (3-4 mm in diameter each) for 48 hours. Then the fiber was glued to a metal table with silicate glue with aluminum shavings. After fixing the fiber on the table, gold is deposited, the thickness of the deposited layer is 15–20 µm. The work was carried out by Jeol JSM 35C and Jeol JSM 6060LA. Preparations for research in scanning electron microscopes Jeol JSM 35C and Jeol JSM 6060LA were attached with epoxy glue with aluminum chips to a stage, kept for three days in a sealed vessel with silica gel cubes and sprayed with gold 15–20 A.

Figure 5: Contacts between representatives of different taxa of microorganisms. Rhizosphere zone of Arabidopsis. Scanning electron microscopy.

Figure 6: Artificial substrate for growing plants.

The substrate for growing plants and researching microorganisms was made from flexible, heat-resistant polyethylene fibers arranged parallel to each other. The preparation of the substrate was carried out as follows: from the fibers of a polymer, non-toxic for living objects, 0.5 mm in diameter, beams with a diameter of 40 mm and a height of 30 mm were formed. The beams were fixed with a metal ring and sealed on one of the transverse sections by heating the metal mold, in which the end of the beam was placed, to the melting point of polyethylene. After the preparation of the substrate, it was processed to eliminate possible toxic impurities and mechanical particles. For this, the items were washed in running water, then boiled for 10 minutes in a 2% CaCO₃ solution, after which they were washed and boiled 3-4 times for 10 minutes in distilled water. The technology of cultivating Arabidopsis plants on an artificial substrate has been worked out. On the new substrate, the full life cycle of Arabidopsis plants was obtained, and viable seeds were obtained. The life cycle averaged 60 days. The obtained images of microbial landscapes, made by the methods of both light and electron microscopy, showed a fundamental possibility. The dynamics of the formation of the substrate microcenosis was also monitored. It was noted that in the early stages of cenosis formation, a large amount of algae is observed in the artificial substrate, especially at the upper edge of the substrate (Figure 2). However, as the plant grows, the amount of algae decreases and bacterial-fungal associations become dominant (Figure 3, 4). The new technology also made it possible to record the intercellular contacts of the cenosis participants using scanning electron microscopy and transmission microscopy (Figure 5). We tried, based on modern possibilities, to create the technology necessary for studying the spatial architecture of the microbial substrate. They began compiling a list of those properties that such a technology should have (a kind of analogue of a "technical design assignment"). This list boiled down to the following:

• The basis of the technology should be a special substrate for growing plants and establishing in it an appropriate rhizosphere microcenosis;
• The substrate should not have cracks, cavities, and its entire surface should be accessible for study;
• The substrate, being the material basis for both plant growth and the formation of microcenoses, should provide in all ranges of space the possibility of carrying out all informative research methods that exist for the study of microorganisms.

In accordance with the formulated properties that such a substrate should have, it was created. In general form, plastics have the widest possibilities for any manipulation. They were taken as the basis for the choice of the substrate material. The organization of the substrate so that it is all available for analysis and, moreover, ensures the safety of the architecture of the microcenosis in its entire range, is possible only in the form of elements oriented in space. They must also provide the necessary capillarity to ensure the water nutrition of plants. At the same time, the biological neutrality of the substrate and its resistance to the degrading effects of both plants and microorganisms must be ensured. Structurally, this was solved in the form of an oriented bundle of polyethylene fibers (in one of the variants), fused from the end into a polyethylene plate (Fig. 6). Capillarity was created in the beam, providing both water and gas regimes, which could be controlled at will. The smooth surface of the fibers and their continuous (without pores, cracks, etc.) cast structure ensured full accessibility to all areas of the substrate (Fig. 7). Neither microflora nor roots, overgrowing fibers at such a surface, cannot escape from direct control. The composition is plastic (in this case, polyethylene, although it could be any other) made it possible to combine all research methods, since, unlike glass, vermiculite, soil silicates, etc., it could be cut on a microtome and examined in an electron microscope. Finally, the continuity of the bundle filaments over the entire height of the substrate made it possible to study the architecture of microcenoses throughout their entire volume both in the transverse and in the longitudinal organization (patent No. 11489; IPC 7 A01G31 / 02; publ. Bull. No. 12, 15.12.2005). Further work was carried out in terms of studying the features and capabilities of the created substrate. Arabidopsis, grown under the conditions of a laboratory analogue of a greenhouse, was chosen as a model plant. Natural substrates (soils) have their own equilibrium-stable microflora (formed cenoses) (Buckley at al., 2003, Gregory, 2006). Artificial, and even those in which there is nothing like this, have a random (and extremely poor) composition of microorganisms. Therefore, if we start from scratch, that is, from a pure bundle of fibers, then the succession begins from some zero mark. The first cultivation of such plants as Arabidopsis usually "does not go". Good growth (Fig. 8) occurs only after several generations of the plant (and even then incomplete - at first they do not reach the end). However, you can use, recreate in the proposed substrate ready-made cenoses (moreover, any of them). For this, the fibers are located in that natural (or artificial) substrate with the formed microflora, which, as the initial one, they want to reproduce. Microflora overgrows the substrate, and then in the already formed system, plants are grown. In this case, the succession will still take place, since the base is substrate. Merely it will start with the existing one, which will become the basis. Full accessibility of the substrate and all components of the cenosis in the entire scope of possibilities - from "talker" to untouched architecture, which allows the study and control with exhaustive completeness to the entire depth of the substrate (from the surface in contact with the atmosphere to the bottom) and to the entire theoretically possible scale magnifications, namely using electronic optics.

Figure 7: Fiber - a constituent element of an artificial substrate for growing plants.

Figure. 8: Growth of Arabidopsis thalian a plant in growing substrate.

Substrates of this type can become a promising basis for artificial systems, since they provide full resolution - assessment, study and control by all available methods of analysis. In addition to a completely artificial substrate, this technology fundamentally makes it possible to study the structure of cenoses in natural conditions. A bundle of any size and length can be made of fibers, including for one ("one piece") plant. The seed sown into it, with preliminary or simultaneous placement of the bundle in the natural substrate, will ensure complete penetration between the fibers of all components of such a substrate (from solutions and colloids to microflora). The plant will grow practically in natural conditions, and the study can be carried out to the full depth under the full resolution conditions provided by the artificial substrate. Due to the fact that such an artificial substrate - "quasi-substrate" can be located, fit, penetrate into the natural substrate, it becomes its integral part. The fragment of the microcenosis formed on is full-sized, and after the removal of the “quasi-substrate-object” of studying what has formed on it, the entire microcenosis in its intact architecture becomes fully accessible to the entire desired depth (thickness) of the substrate with a gradient of conditions. The very organization of the placement of the "quasi-substrate-object" provides full accessibility for any study in any size in terms of the area and volume of the substrate. The coenosis in artificial substrates must be controlled. Only in substrates of full resolution this is fundamentally possible due to exhaustive control over the state of cenoses and their dynamics. This becomes possible by changing the conditions, if necessary, then generally introducing some additional, pre-prepared biological components in accordance with the information obtained by monitoring the state of the cenosis, and if they deviate from the desired, making changes to the cultivation mode and tracking the changes in cenosis occurring under their influence.

1. Copley J, Ecology goes Nature. 2000; 406: 452-454.
2. Buckley DH, Schmidt TM, Diversity and dynamics of microbial communities in soils from agro-ecosystems. Environmental Microbiology. 2003; 5: 441-452.
3. Buckley DH, Schmidt TM, The structure of microbial communities in soil and the lasting impact of Microbial Ecology. 2001; 42: 11-21.
4. Akkermans ADL, Mirza MS, Harmsen HJM, Blok HJ, Herron PR, Sessitsch A, et al. Molecular ecology of microbes A review of promises, pitfalls and true FEMS Microbiology Reviews. 1994; 15: 185-194.
5. El-Shatnawi MKJ, Makhadmeh IM, Ecology of the plant-rhizosphere. 2001.
6. Gregory PJ, Roots, rhizosphere and soil: the route to a better understanding of soil science? European Journal of Soil Science. 2006; 57: 2-12.
7. Bartoli F, Gomendy VG, Royer JJ, Niquet SO, Vivier H, Grayson RA, et al. multiscale study of silty soil European J of Soil Sci. 2005; 56: 207-223.
8. Hinsinger P, Gobran GR, Gregory PJ, Wenzel WW, Rhizosphere geometry and heterogeneity arising from rootmediated physical and chemical processes. New Phytologist. 2005; 168: 293-303.
9. Greaver TL. Sternberg LSL. Fluctuating deposition of ocean water drives plant function on coastal sand dunes. Global Change 2007; 13: 216-223.
10. Amann RI, Ludwig W, Schleifer KH, Phylogenetic identification and in situ detec- tion of individual microbial cells without cultivation. Microbiological Reviews. 1995; 59:143-169.

11. Vinogradsky S.N. Soil microbiology. Ministry of Education and Science of the USSR Academy of Sciences, 1952-792.

12. Methods for studying soil microorganisms and their metabolites Ed. N. A. Krasilnikova. M. MSU, 1966; 216

13. El?Shatnawi MKJ, Makhadmeh MI. System. Review. Agronomy and Crop Science, 187: 1-9.
14. Joseph SJ, Hugenholtz P, Sangwan P, Osborne CA, Janssen1 PH. Laboratory cultivation of widespread and previously uncultured soil bacteria. Applied and Environmental Microbiology. 2003; 69: 7210-7215.