Open access

Planar Oxygen Sensors for Non Invasive Imaging in Experimental Biology

Written By

Henning Tschiersch, Gregor Liebsch, Achim Stangelmayer, Ljudmilla Borisjuk and Hardy Rolletschek

Submitted: October 23rd, 2010 Published: June 9th, 2011

DOI: 10.5772/17893

Chapter metrics overview

3,377 Chapter Downloads

View Full Metrics

1. Introduction

The presence of molecular oxygen is a sine qua non for aerobic metabolism. In both plant and animal mitochondria, it acts as the terminal electron acceptor for oxidative phosphorylation occurring during cellular respiration, and is necessary for the generation of ATP, the common energy currency within the living cell (Atkinson, 1977; Cooper, 2000). It is the major by-product of photosynthesis in which plant biomass is accumulated by the conversion of carbon dioxide into polymeric compounds. Since respiration and photosynthesis are so fundamental to life on earth, an understanding of the mechanisms underlying oxygen consumption, production and homoeostasis has become a significant field of both biological and biotechnological research (Volkmer et al., 2009).

Oxygen (micro-) sensors, which are widely used in the life sciences, are designed to provide a precise measurement of the concentration of oxygen within a localized region of a tissue or an organ (Borisjuk & Rolletschek, 2009 ). Most of these devices have been based on miniaturized Clark-type electrodes (Revsbech & Jørgensen, 1986), in which oxygen diffuses into the sensor via a permeable membrane, following which its reduction at the cathode generates a measurable electrical current. This approach can deliver a spatial resolution at the low µm scale. Increasingly this technology is being replaced by optical oxygen microsensors (micro-optodes) based on fibre optic materials (Klimant et al., 1995; Rolletschek et al., 2009), in which the concentration is assessed in tapered glass fibres of tip size ~50µm via the dynamic quenching of a luminophore. This approach enjoys several advantages over the electrochemical detection system, as detailed elsewhere (Kühl & Polerecky, 2008; Rolletschek et al., 2009).

Importantly, microsensor-based approaches are invasive, which means that a given biological sample cannot be readily studied over a prolonged time period. Furthermore, the internal structure of most biological samples is far from homogeneous, with complex compartmentation being the norm. As a result, whole tissue measurements can only reflect the mean performance of a tissue, and cannot report variation between distinct compartments. This loss of richness compromises the value of such data for elucidating the biology of the tissue as a whole. At best, conventional sensor systems assess oxygen concentrations across a transect, leaving its two dimensional distribution unknown. Lifting this limitation requires the development of a planar sensor.

Here, we present a novel oxygen sensing approach, in which image processing has been combined with optical sensor technology. The optical sensor foil (i.e. the planar optode) attached to the surface of the sample translates the oxygen signal into a light signal, which is then captured and interpreted pixel by pixel by a digital camera. Since a single image captures an array of sensor points, the system permits an instantaneous two-dimensional mapping of oxygen distribution. While some analagous approaches have already been described in the literature (Liebsch et al., 2000; Glud et al., 2005; Kühl & Polerecky, 2008), the system we describe here represents a significant improvement with respect to spatial resolution, handling and image processing, and eventually ease of use. Two applications of the system are described in some detail: the first involves a respiring (oxygen consuming) root of oilseed rape (Brassica napus), and the second a photosynthetically active (oxygen generating) leaf of Cabomba caroliniana, an aquatic perennial herbaceous plant. In both, marked oxygen gradients were detected across both time and space. In combination with the use of specific inhibitors, the planar sensor system can be expected to permit a spatially well resolved analysis of respiration or photosynthesis. We conclude that the new planar sensor setup provides fascinating opportunities for research in all areas of life sciences.


2. Planar oxygen sensors – design, calibration and applications

The following chapters provide an overview on (i) the technical features of the novel planar sensor setup, and (ii) the possibilities for its use in plant biology, in particular to study respiration (oxygen consumption) and photosynthesis (oxygen production).

2.1. Experimental design for life time imaging of oxygen

Digital revolution in photography induced a giant trend towards capturing images and creating movies of nearly everything one can think of. Beside scientific and industrial cameras the market of consumer imaging devices is constantly growing and continually new products are launched showing increased resolution while being miniaturized. The enhancement of image quality and downsizing affects all market segments of consumer cameras, high-quality SLR cameras as well as low-tech webcams and mobile phone cameras. As a result, the use of such consumer devices is also of increasing interest in the field of opto-chemical sensing where the response of a fluorescent sensor is recorded in order to measure chemical analytes. Typically, for this application fairly bulky and sophisticated camera systems (Holst et al., 1998; Schröder et al., 2007; Kühl & Polerecky, 2008) are used which support time resolved measurement. Measuring a lifetime dependent parameter is generally preferred because of the favourable accuracy due to suppressing common interferences including heterogeneous lightfield or sample coloration and auto-fluorescence allowing even transparent sensor foils (Holst et al., 2001). This is not possible if using even high-tech standard consumer cameras which allow ratiometric calibration schemes at best. However, beside the restriction of transparent sensors ratiometric imaging has proved to be also an excellent solution for measuring analyte contents of a sample quantitatively and two dimensionally (Wang et al., 2008). Then, it depends on the sample target and analytical problem if the possibility of miniaturisation and mobility overcompensates the restriction. Especially in biological application fields of imaging with fluorescent optical sensors it is desirable to use compact devices which are close to pocket size and can easily be taken to the place of measurement. As a result, complex biological systems are not disturbed and can be measured “as is” in their natural environment or green house. New reports address the topic of applying portable consumer technology by using SLR cameras (Wang et al., 2010) or even mobile phones (Filippini & Lundstrom, 2006) with the side-effect of substantially reducing the costs for imaging devices at the same time. However, in these solutions the question about suitable optical filters, macro lense and light source combinations is not sufficiently solved. Especially a micrometer resolution is indispensable if investigating biological processes of plant seeds, embryos, collenchyma or rhizospheres. Also in other medical and biotechnical applications a high resolution is needed including monitoring phase transitions in aquatic biology, mini bioreactors, tissue engineering and skin microcirculation. Therefore, we developed the idea of using consumer camera technology further and identified a type which fits perfectly to the demands of fluorescent optical imaging: the USB microscope. The results presented here were measured with a prototype of new imaging product series “VisiSens” (PreSens GmbH, Regensburg, Germany).

The market of USB microscopes is allocated to many types showing huge differences in image quality. We based our development on a current high-end USB microscope with good sensitivity and image quality and improved it for imaging fluorescence-optical sensors by integrating an optic block with high quality LED PCB and optical filters. Figure 1 shows a solid (left panel) and a transparent (middle panel) technical drawing of the measurement head, showing its compactness and the arrangement of the respective components. The three images beside show millimeter paper measured with different magnification settings. Maximum magnification is approximately 200-fold where the field of view is ~ 2.5 x 2.0 mm.

Figure 1.

a) Solid and (b) transparent technical drawing of the compact measurement head incorporating a USB-microscope for imaging fluorescent sensor foils. Camera and light source are powered via standard USB connector. (c) Images of millimeter paper demonstrating the magnification up to 200-fold.

Figure 2 shows an explosion drawing of the USB microscope where the components are addressed in detail. The all-aluminium detector head (a) integrates a color RGB CMOS chip (b), a microscope lense (c) with manual focus, 8 blue emitting LEDs (i) which are driven by a printed circuit board (PCB) (g) and aligned in an aluminium block (h) and optical filters for light diffusion (j), excitation (k) and emission (l). The up to 200-fold magnified images are recorded with a 1.3 megapixel (1280 x 1024) color chip which results in more than 300,000 independent sensing points (= pixel) for the respective sensor response (i.e. color channel of the RGB chip). Maximum spatial image resolution is ~ 2.5 mm per 1280 pixel (~ 2 µm per pixel). Maximum spatial sensor resolution depends on the sensor used and is typically ~ 25 to 100 µm. Power supply of the camera and the LED light source is provided via the standard USB connector which makes the system laptop compatible and a portable device. The dimensions of the detection head are 10 cm in length and 3.8 cm in diameter, the working distance is typically from 1 to 5 cm. The camera can be used free hand or mounted to a stand.

Figure 2.

Explosion drawing of the PreSens USB-microscope “VisiSens” optimized for imaging of fluorescencent sensor foils: (a) aluminium housing, (b) RGB CMOS-chip, (c) opening for manual focus, (d) microscope lense, (e) filter tube, (f) mounting and adjustment ring(g) transmitter PCB, (h) LED reflector block, (i) LEDs, (j) diffusor, (k) short pass excitation filter, (l) long pass emission filter and (m) sapphire glass.

For the measurement the imaging system uses flexible sensor foils which allow two dimensional recording of oxygen distributions in aqueous phase over an area typically ranging from 5 x 5 mm2 up to 40 x 40 mm2. The planar sensors consist of an oxygen sensitive dye and a reference dye which are immobilized in an oxygen permeable polymer matrix and fixed on a transparent polyester support and overlaid with a white oxygen transparent layer for optical isolation. We used a PreSens sensor foil which is not described here but similar to that described in detail by Wang et al., (2010).

During the measurement the sensitive layer is in contact with the sample and the fluorescence is measured from the backside. Every single indicator dye molecule is interacting independently with oxygen in the form that the red fluorescence of the sensitive dye is dynamically quenched. This means that the energy of the excited dye is transferred to the oxygen molecule by collision (see Fig. 3) and consequently the intensity of the sensitive dye is reduced with increasing oxygen content of the sample.

The reference dye, however, is not affected by oxygen and shows constantly a green light signal. The working range of the oxygen sensor covers the typical biological range from 0 to 100% air saturation (corresponding to 6.04 mL of oxygen per liter freshwater at 25º C and 101.3 kPa standard atmosphere). The sensors can be shaped to any desired geometry using a scissor.

Figure 3.

A) Schematic of the sensor foil and principle of dynamic fluorescence quenching. The sensor responds to oxygen by passing the energy of an excited state dye via collision to the oxygen molecule. This results in fluorescence quenching with increasing oxygen content of the sample. (B) Measurement set-up. The sensor consists of an indicator doped polymer layer which is fixed on a transparent polyester support. During the measurement the sensor has to be in direct contact with the sample. The sensor response is measured from outside through a transparent window of the respective vessel.

For quantitative evaluation of the sensor response we applied a ratiometric calibration scheme. The sensitive dye and reference dye are excited with the identical light source at the same time but emit at different bands of wavelength. In our case, both dyes are excited with blue light and while the sensitive dye emits red light, the reference emits green light. We selected dyes whose emission bands of wavelength meet exactly the red and green channel sensitivity of our color RGB chip. This enables to obtain the green reference information independently from the red sensor information within a single image at the same time. A quantitative evaluation is done by rationing the red and green channel of the RGB image in order to reference out the main interferences of intensity based measurements namely inhomogeneous light field and dye concentration including varying sensor layer thickness. The respective oxygen content is computed from the ratio applying a calibration function which was derived from measuring the sensor response at known oxygen concentrations in a chamber (Figure 4).

Figure 4.

A) Calibration Images and data. The sensor was exposed to different oxygen gas mixtures and the response was imaged. (B) The Stern-Vollmer plot leads to a linear relation which was used for calculating the respective oxygen contents of the measurement images.

2.2. Application of planar optical sensors for oxygen measurements on plants

Unlike animals, plants are both producers and consumers of oxygen via photosynthesis and cellular respiration, respectively. Plant leaves, stems and seed tissues generally possess chloroplasts, producing oxygen under illumination ((Borisjuk and Rolletschek, 2009 ; Tschiersch et al., 2011) whereas roots are a typical example of non-green tissues. Their oxygen homoeostasis and exchange capabilities depend on the developmental state (age), several tissue characteristics (e.g. cuticula) as well as environmental conditions (e.g. temperature) (Armstrong et al., 1994, 2009). In particular, the complex anatomy of tissues often hampers oxygen diffusion, thereby causing steep diffusion gradients and local oxygen deficiencies.

Planar oxygen sensors are an alternative to conventional microsensor-based techniques, and allow to viualize the oxygen dynamics between tissues and surrounding media. Here, we demonstrate the applicability of the planar sensor system to study oxygen dynamics in two plant models: (i) the respirating (O2-consuming) root system of the crop plant oilseed rape (Brassica napus), and (ii) the photosynthetically active (O2-producing) leaf of the water plant Cabomba caroliniana.

2.2.1. Imaging of oxygen consumption in living plant roots

Several contrasting plant species, that differ in their relative growth rates (herbs, grasses, shrubs and trees), possess respiration rates in a relative narrow range between 20 and 52 nmol oxygen (g DW)-1 s-1 (Loveys et al., 2003). Similar values were reported for roots of crop plant seedlings using clark-type electrodes (Tab. 1). For roots of oilseed rape seedlings we measured mean respiration rates of ~ 79 nmol oxygen (g DW)-1 s-1 (own unpublished data).

Species Root respiration rate [nmol O2 (gDW)-1 s-1] References
Oilseed rape (Brassica napus) 79.3 this study
Barley (Hordeum vulgare) 16.2 Bloom et al., 1992
Wheat (Triticum aestivum) 64.0 Kurimoto et al., 2004
Rice (Oryza sativa) 39.1 Kurimoto et al., 2004
Corn (Zea mays) 56.7 Hejl & Koster, 2004
Potato (Solanum tuberosum) 14.8 Bouma et al., 1996
Tomato (Solanum lycopersicum) 15.8 Hadas & Okon, 1987
Pea (Pisum sativum) 69.2 DeVisser et al., 1986
Soybean (Glycine max.) 67.5 Millar et al., 1998

Table 1.

Root respiration rates of various crop plants

Using the planar oxygen sensor we here aimed to visualize quantitatively the oxygen consumption of intact roots. For this purpose oilseed rape was grown on 0.9% Difco-agar for 14 days. For the measurement the root segments of seedlings were covered with the transparent sensor foil. The use of transparent foils is of special importance because it allows the alignment between the sample structure and the measured oxygen distribution. The sample was enclosed by an incubation chamber to limit oxygen diffusion from the outside. This custom-made system allowed the combined imaging of the root (Fig. 5a) and the oxygen concentration (Fig. 5b). In our experiment, the oxygen distribution in the sample was followed over six hours with measuring points every 15 min. Based on the decline in oxygen concentration over time, the respiration rate of the central root zone was calculated as 0.015 % air saturation min-1, which corresponds to ~ 12.5 µmol oxygen h-1. The final image of oxygen distribution in Fig. 5b demonstrates that oxygen consumption of the root system can be mapped for distinct root regions with sub-millimetre resolution using the planar sensor setup. It should be noted that the use of thin sensor foils and incubation chambers is preferable, because this reduces diffusive smearing of the spatial oxygen distribution, which otherwise can mask the true oxygen dynamics as well as its gradients. Thus, it allows maximizing the spatial resolution in oxygen mapping.

Figure 5.

Imaging of oxygen consumption for roots of oilseed rape. (A) Picture of the root system; the square indicates the area where oxygen imaging was applied. (B) Image of oxygen distribution after 6 h of dark incubation. (C) Time course of oxygen concentration measured within the main root area (region indicated by the star in (A))

2.2.2. Monitoring of oxygen dynamics in photosynthetically-active leaves

The water plant Cabomba caroliniana is known as a classical object in photosynthesis research (Smith, 1936; Kok, 1949). Traditionally, fronds of Cabomba were used in the classroom to demonstrate bubble formation (photosynthetic oxygen evolution) upon illumination. Cabomba leaves freshly collected from surface waters show a photosynthetic rate at light saturation of about 2.5 µmol CO2 consumed (mg Chl)-1 h-1 and a respiratory rate of 2-3 µmol CO2 evolved (mg Chl)-1 h-1 (Van et al., 1976). In a more recent study, respiration rates of up to 30 µmol O2 (g DW)-1 h-1 were reported (Penuelas et al., 1988).

Here, we aimed to visualize both consumption and production of oxygen in the Cabomba leaf. For this purpose, one Cabomba frond was covered with the sensor foil, and enclosed in the incubation chamber (restricting oxygen diffusion from the exterior). This custom-made system allowed the combined imaging of leaf structure and oxygen concentration. Figure 6a shows a picture of the Cabomba leaf. Its incubation under non-lit conditions for only 30 min led to a remarkable decrease in oxygen concentration, which was attributable to cellular respiration. The overlay of the color image and the sensor response image in Fig. 6b demonstrates that this decrease co-localised with the characteristic leaf shape of the plant. The oxygen level declined to ~13% of air saturation after 60 min under non-lit conditions (Fig. 6c). This indicates a substantial respiratory activity of Cabomba leaves. Upon illumination (15 µmol photons m-2 s-1), a quick increase in oxygen concentration became apparent (Fig. 6c), indicating a strong oxygen production by the leaf. This activity could again be clearly aligned to the leaf shape (Fig. 6d versus 6e).

We suggest that the increase in oxygen concentration under lit conditions was caused by leaf photosynthesis. To prove this suggestion, we performed a similar light/dark experiment with Cabomba leaves in the presence of the photosynthesis inhibitor 3-(3,4-Dichlorophenyl)-1,1-dimethylurea (DCMU). The addition of 20µM DCMU completely abolished the light-dependent oxygen production of the leaf (Fig. 6c). This clearly demonstrates that the light-induced increase in oxygen levels is due to photosynthesis.

The differences between the oxygen exchange rates in darkness and light were remarkable, and demonstrate the high photosynthetic capacity of Cabomba caroliniana even under low light supply. The imaging experiment revealed that (i) the oxygen production under low light (15 µmol photons m-2 s-1) fully compensated dark respiration, and (ii) the oxygen evolution rate in the light was five times higher as compared to the oxygen consumption rate in the dark. This result was confirmed by conventional microsensor-based analysis (optical oxygen sensor SP-PSt3 connected to Fibox3 oxygen meter; PreSens Sensing GmbH, Germany). Under non-lit conditions, a respiration rate of 26.1 µmol O2 (g DW)-1 h-1 was measured for Cabomba leaves. Under lit conditions (< 10 µmol photons m-2 s-1), the photosynthetic oxygen production was sufficient to fully compensate oxygen consumption by respiration (own unpublished data).

The planar sensor system can be used to monitor fast responses of leaves to dark/light switches. Figure 7 displays in detail the differential effect of light versus darkness on the oxygen exchange of the Cabomba leaf. Within a few minutes clear changes in oxygen concentration could be monitored in darkness. Extended incubation under dark resulted on the one hand in a stronger decrease in oxygen concentration and on the other hand in a smearing of the sensor signal due to the diffusion of oxygen (Fig. 7, left panels). Upon start of illumination (15 µmol photons m-2 s-1), the fluorescence signal declines steadily, which indicates the photosynthetic oxygen release. We conclude that the planar sensor system can perform both spatial and temporal oxygen monitoring.

Figure 6.

Imaging of oxygen exchange by the leaf of Cabomba caroliniana. (A) Color image of the leaf. (B) Superimposed images of leaf and oxygen distribution after 20 min dark incubation. (C) Time course of oxygen dynamics of leaf with and without addition of DCMU; analysed region corresponds to 1.4 mm2 leaf area marked with a star in (A). (D) Image of oxygen distribution after 30 min dark incubation. (E) Image of oxygen distribution after 30 min light incubation (15 µmol photons m-2 s-1) subsequent to dark incubation.

Figure 7.

Monitoring of oxygen dynamics of a Cabomba caroliniana leaf during 15 min of dark incubation and the subsequent light phase (15 µmol photons m-2 s-1).


3. Prospects for planar oxygen sensing

Above data clearly illustrate the major advantage of planar oxygen sensing as a non-invasive imaging technique. For the first time, rates of oxygen production and consumption could be spatially resolved and visualized. The acquired color-coded oxygen maps are quantitative and have a resolution in the sub-millimetre range. In this way, dynamic changes in oxygen concentration within the complex root (leaf) system of a plant and its surrounding media can be studied. This non-invasive approach will allow investigating mechanisms of cellular growth and interactions among organisms and their environment. Although not studied here, the planar oxygen sensor system will be of high significance in other research areas like biotechnology or medicine. For example, documenting the oxygen dynamics during cell infection and cancerogenesis could help identify specific drug targets to slow or stop the uncontrolled growth of cancer cells (Babilas et al., 2005).

Currently, planar sensors have been developed to specifically detect oxygen, carbon dioxide or pH. It is also conceivable that planar sensor foils have multi-analyte properties. These sensors will combine an oxygen sensitive dye (and its reference dye) with dyes specific for other analytes. In this way, several analytes can be quantitatively visualized in single experiments.



We acknowledge funding by the Bundesministerium für Wirtschaft und Technologie within the framework of Zentrales Innovationsprogramm Mittelstand (ZIM). We also wish to thank Steffen Wagner for excellent technical assistance.


  1. 1. Armstrong W. Strange M. E. Cringle S. Beckett P. M. 1994 Microelectrode and modelling study of oxygen distribution in roots. Annals of Botany, 74 3 (September 1994), 287 299 , 0305-7364
  2. 2. Armstrong W. Webb T. Darwent M. Beckett P. M. 2009 Measuring and interpreting respiratory critical oxygen pressures in roots. Annals of Botany, 103 2 (January 1994), 281 293 , 0305-7364
  3. 3. Atkinson D. E. (ed. 1977 Cellular energy metabolism and its regulation. Academic Press, 0-12066-150-0 York, USA
  4. 4. Babilas P. Liebsch G. Schacht V. Klimant I. Wolfbeis O. S. Szeimies R. M. Abels C. 2005 In vivo phosphorescence imaging of pO2 using planar oxygen sensors. Microcirculation, 12 6 (September 2005), 477 487 , 1073-9688
  5. 5. Bloom A. J. Sukrapanna S. S. Warner R. L. 1992 Root respiration associated with ammonium ore nitrate absorption and assimilation by barley. Plant Physiology, 99 4 (August 1992), 1294 1301 , 0032-0889
  6. 6. Borisjuk L. Rolletschek H. 2009 The oxygen status of the developing seed. New Phytologist, 182 1 (April 2009), 17 30 , 0002-8646X
  7. 7. Bouma T. J. Broekhuysen A. G. M. Veen B. W. 1996 Analysis of root respiration of Solanum tuberosum as related to growth, ion uptake and maintenance of biomass. Plant Physiology and Biochemistry, 34 6 (June 1996), 795 806 , 0981-9428
  8. 8. Cooper G. M. 2000 The Cell- A Molecular Approach (2nd edition), Sinauer Associates, 0-87893-106-6Sunderland (MA),USA
  9. 9. De Visser R. Brouwer K. S. Posthuma F. 1986 Alternative path mediated ATP-synthesis in roots of Pisum sativum upon nitrogen supply. Plant Physiology, 80 2 (February 1986), 295 300 , 0032-0889
  10. 10. Filippini D. Lundstrom I. 2006 Method and system for chemical or biochemical analysis of a target analyte in a target environment. US Pat. 7,092,089.
  11. 11. Glud R. N. Wenzhöfer F. Tengberg A. Middelboe M. Oguri K. Kitazato H. 2005 Distribution of oxygen in surface sediments from central Sagami Bay, Japan: In situ measurements by microelectrodes and planar optodes. Deep Sea Research Part I: Oceanographic Research Papers, 52 10 (October 2005), 1974 1987 , 0967-0637
  12. 12. Hadas R. Okon Y. 1987 Effect of Azospirillum brasilense inoculation on root morphology and respiration in tomato seedlings. Biology and Fertility of Soils, 5 3 (December 1987), 241 247 , 0178-2762
  13. 13. Hejl A. M. Koster K. L. 2004 Juglone disrupts root plasma membrane H+-ATPase activity and impairs water uptake, root respiration and growth in soybean (Glycine max.) and corn (Zea mays). Journal of Chemical Ecology, 30 2 (February 2004), 453 471 , 0098-0331
  14. 14. Holst G. Grunwald B. 2001 Luminescence lifetime imaging with transparent oxygen optodes. Sensors and Actuators B, 74 1-3 , (April 2001), 78 90 , 0925-4005
  15. 15. Holst G. Kohls O. Klimant I. König B. Kühl M. Richter T. 1998 A modular luminescence lifetime imaging system for mapping oxygen distribution in biological samples. Sens. Actuators B, 51 1-3 , (August 1998), 163 170 , 0925-4005
  16. 16. Iimant K. I. Meyer V. Kühl M. 1995 Fiber-optic oxygen microsensors, a new tool in aquatic biology. Limnology and Oceanography, 40 6 (May 1995), 1159 1165 , 1541-5856
  17. 17. Kok B. 1949 On the interrelation of respiration and photosynthesis in green plants. Biochimica et Biophysica Acta, 3 1 (January 1949), 625 631 , 0005-2728
  18. 18. Kühl M. Polerecky L. 2008 Functional and structural imaging of phototrophic microbial communities and symbioses. Aquatic Microbial Ecology, 53 1 (September 2008), 99 118 , 0948-3055
  19. 19. Kurimoto K. Day D. A. Lambers H. Noguchi K. 2004 Effect of respiratory homeostasis on plant growth in cultivars of wheat and rice. KurimotoK.DayD. A.LambersH.NoguchiK. (2004). Effect of respiratory homeostasis on plant growth in cultivars of wheat and rice. Plant Cell and Environment, Vol.27, No.7, (July 2004), pp. 853-862, ISSN 0140-7791 Cell and Environment, 27 7 (July 2004), 853 862 , 0140-7791
  20. 20. Liebsch G. Klimant I. Frank B. Holst G. Wolfbeis O. S. 2000 Luminescence lifetime imaging of oxygen, pH, and carbon dioxide distribution using optical sensors. Applied Spectroscopy, 54 4 (April 2000), 548 559 , 0003-7028
  21. 21. Loveys B. R. Atkinson L. J. Sherlock D. J. Roberts R. L. Fitter A. H. Atkin O. K. 2003 Thermal acclimation of leaf and root respiration: an investigation comparing inherently fast- and slow- growing plant species. Global Change Biology, 9 6 (June 2003), 895 910 , 1365-2486
  22. 22. Millar A. H. Atkin O. K. Menz R. I. Henry B. Faquhar G. Day D. A. 1998 Analysis of respiratory chain regulation in roots of soybean seedlings. Plant Physiology, 117 3 (July 1998), 1083 1093 , 0032-0889
  23. 23. Penuelas J. Murillo J. Azcon-Bieto J. 1988 Actual and potential dark respiration rates and different electron transport pathways in freshwater aquatic plants. Aquatic Botany, 30 4 (May 1988), 353 362 , 0304-3770
  24. 24. Revsbech N. P. Jørgensen B. B. 1986 Microelectrodes: their use in microbial ecology. In: Advances in Microbial Ecology, 9 K.C. Marshall (ed.), 293 352 , Springer, 0-30642-184-4 York, USA
  25. 25. Rolletschek H. Stangelmayer A. Borisjuk L. 2009 The methodology and significance of microsensor-based oxygen mapping in plant seeds- an overview. Sensors, 9 5 (April 2009), 3218 3227 , 1424-8220
  26. 26. Schröder C. R. Neurauter G. Klimant I. 2007 Luminescent dual sensor for time-resolved imaging of p CO2 and p O2 in aquatic systems. Microchimica Acta, 158 3-4 , (May 2007), 205 218 , 0026-3672
  27. 27. Smith E. L. 1937 The influence of light and carbon dioxide on photosynthesis. The Journal of General Physiology., 20 6 (July 1937), 807 830 , 0022-1295
  28. 28. Tschiersch H. Borisjuk L. Rutten T. Rolletschek H. 2011 Gradients of seed photosynthesis and its role for oxygen balancing. Biosystems, 103 2 (February 2011), 302 308 , 0303-2647
  29. 29. Van T. K. Haller W. T. Bowes G. 1976 Comparison of photosynthetic characteristics of three submersed aquatic plants. Plant Physiology, 58 6 (December 1976), 761 768 , 0032-0889
  30. 30. Volkmer E. Drosse I. Otto S. Stangelmayer A. Stengele M. Cherian Kallukalam. B. Mutschler W. 2008 Hypoxia in static and dynamic 3D culture systems for tissue engineering of bone. VolkmerE.DrosseI.OttoS.StangelmayerA.StengeleM.CherianKallukalam. B.MutschlerW. (2008). Hypoxia in static and dynamic 3D culture systems for tissue engineering of bone. Tissue Engineering Part A, Vol.14, No.8, (August 2008), pp. 1331-1340, ISSN 2152-4947 Part A, 14 8 (August 2008), 1331 1340 , 2152-4947
  31. 31. Wang X. D. Chen X. Xie Z. X. Wang X. R. 2008 Reversible optical sensor strip for oxygen. Angewandte Chemie, 120 39 (September 2008), 7560 7563 , 1521-3757
  32. 32. Wang X. D. Meier R. J. Link M. Wolfbeis O. S. 2010 Photographing oxygen distribution. Angewandte Chemie International Edition, 49 29 (July 2010), 4907 4909 , 1433-7851

Written By

Henning Tschiersch, Gregor Liebsch, Achim Stangelmayer, Ljudmilla Borisjuk and Hardy Rolletschek

Submitted: October 23rd, 2010 Published: June 9th, 2011