COMPARING PERFORMANCE OF CROP SPECIES MIXTURES AND PURE STANDS

Highlights

•The literature on intercropping comprises thousands of papers.

•Evidence synthesis is needed to develop general conclusions.

•Quantitative evidence synthesis requires meaningful comparative performance metrics.

•The background, meaning, and limitations of some performance metrics is explained.

•Future challenges are identified.

Abstract

Intercropping is the planned cultivation of species mixtures on agricultural land. Intercropping has many attributes that make it attractive for developing a more sustainable agriculture, such as high yield, high resource use efficiency, lower input requirements, natural suppression of pests, pathogens and weeds, and building a soil with more organic carbon and nitrogen. Information is needed which species combinations perform best under different circumstances and which management is suitable to bring out the best from intercropping in a given production situation. The literature is replete with case studies on intercropping from across the globe, but evidence synthesis is needed to make this information accessible. Meta-analysis requires a careful choice of metric that is appropriate for answering the question at hand, and which lends itself for a robust meta-analysis. This paper reviews some metrics that may be used in the quantitative synthesis of literature data on intercropping.

Graphical abstract

Keywords

intercropping / species mixtures / meta-analysis / metrics / indicators

1 INTRODUCTION

Interactions between above and below ground structures involve energy, matter and information^[1]. Occam’s razor implies that the models initially considered should only involve energy and matter in determining allocation of resources between above and below ground structures. Such models can go some way to explaining the observed allocation of resources, for example, to water, carbon dioxide, light, and major nutrients acquisition, and the response to defoliation and root pruning^[2–9]. However, such models are less successful when applied to minor nutrients, nutrient toxicity and temperature variations, and to reproduction and storage^[4]. There are also differential above and below ground responses to competition^[10]. Accordingly, as well as energy and matter in above and below ground interactions, it is also necessary to consider the flow of information.

2 ENERGY

Perhaps the most obvious means of transmitting energy from above ground to below ground structures is as photosynthate, i.e., organic carbon such as sucrose, sugar alcohols and/or oligosaccharides, moving down the phloem, thereby allowing catabolism below ground reducing NAD⁺ or NADP⁺ and phosphorylating ADP. Also, the driving force for Münch pressure flow from shoot to root is ATP-consuming proton ATPase, and sucrose-proton symporter in the companion cell-sieve tube element plasma membrane.

Energy is also transmitted via the tension generated by transpiratory water loss^[11,12]. The latent heat of volatilization of water is supplied by photons absorbed by the photosynthetic structure and not stored in the products of CO₂, NO₃^–, SO₄^2– or O₂ photoreduction, fluoresced, or lost as longwave radiation. The liquid water that has been transpired is, in the steady-state, replaced by water moving up the xylem and, ultimately, from the root medium. The movement of water up the xylem when transpiration occurs, and in the absence of root pressure, is driven by water loss and transmitted by tension in the water column, dependent on the rigidity of the walls of xylem conduits that prevents collapse of the conduits, and the absence of pores in the cell wall large enough to allow air from intracellular spaces to enter the xylem under the observed pressure difference between the gas space and the water under tension. In the root system, the negative pressure in the xylem (less negative than in the shoot) drives water flux to the xylem from the soil solution. In addition to the water lost in transpiration, tension in the xylem conduit driven by transpiration supplies water used in cell expansion, as a substrate for photosynthesis, and water cycling down the phloem in Münch pressure flow^[13,14].

Under conditions of limited or no transpiration, water can be moved up the xylem by root pressure, supporting cell expansion, guttation and water cycling down the phloem in Münch pressure flow especially in the case of nutrient transport to the shoot when nutrients are only supplied in the daily dark period^[13]. The widely accepted mechanism of root pressure is the accumulation of solutes in the xylem conduits, decreasing the water potential in the xylem sap relative to that in the root medium. There is some evidence that a component of root pressure can, in some cases, be active water transport, i.e., energized movement of water against a water potential difference^[15].

In addition to tension-driven flow in xylem, and pressure-driven flow in phloem and xylem (root pressure), there are other possibilities of energy transfer from above ground to below ground structures. One is transmission of light along plant axes from illuminated shoots to darkened roots, i.e., light piping, Light transmission along shoot and root axes has been for woody plants^[16] and for herbaceous plants^[17]. Wavelengths usable in photosynthesis were attenuated by about an order of magnitude over a path of 10 mm, with rather less attenuation of 730 nm radiation absorbed by the far-red form of phytochrome^[16,17]. However, energetic use of light in photosynthesis is less likely, granted attenuation of radiation by an order of magnitude per cm: by 30 mm the noon incident 40–700 nm photon flux density of ~2 mmol·m⁻²·s⁻¹ is decreased to ~2 μmol·m⁻²·s⁻¹, less than the light compensation point for net photosynthesis in vascular plants. Below ground structures of terrestrial plants are not photosynthetic, but roots of dark-grown Zea mays seedlings contain protochlorophyllide that can be converted to chlorophyll by red light, although this was applied to the whole seedlings^[18]. A further possibility of energy transfer between above and below ground structures is transmission of electrochemical potential differences, typically as proton motive force, along membranes between photochemical, redox reaction or ATPase-driven primary active transport at one location and the sites of proton motive force use in cotransport^[19]. However, this transmission is unlikely to occur over more than a few 100 μm at most^[19]. As with light, axial transmission of information by electrochemical potential differences by ions in action potentials or variation potentials, for example, along phloem, can occur over much long distances than transmission of proton motive force.

3 MATTER: XYLEM AND PHLOEM TRANSPORT OF WATER AND SOLUTES

The apoplasmic xylem conduits transport soil-derived water and solutes from the below ground to the above ground parts of vascular land plants; acquisition of nutrients from the above ground environment is minimal in most natural environments^[20,21]. The driving force for most of the xylem flux is transpiratory water loss ultimately powered by solar radiation, directly through absorption by photosynthetic structures, and indirectly via wind. This transpiratory loss is much smaller in the dark in C₃ and C₄ plants even when stomata are open. Plants expressing Crassulacean acid metabolism have the highest stomatal conductance at night when dark CO₂ assimilation occurs. Transpiration-driven xylem flux involves negative pressure in the conduits. In contrast, root pressure, involving higher solute concentrations in the xylem conduits than in the root medium and, arguably, with active water transport, involves much slower solution movement in the xylem than in transpiratory water flux in the light, albeit with a higher solute concentration. The apoplasmic nature of the transport pathway imposes fewer constraints on the solute composition than in the symplasmic phloem, especially since the total solute concentration in the xylem is less than that in the phloem. Most of the water transported up the xylem is transpired; the rest is used in the hydration of growing cells, and less is used as the source of electrons in photosynthesis with production of O₂. The universal solute transport role of xylem is movement of solutes derived from the soil to transpirational termini with, for some solutes, transfer to the phloem for transport to growing above ground tissues with limited transpiratory water loss^[22]. As will be seen below, some of these solutes, particularly NH₄⁺, N₂ and sometimes NO₃^–, are subject to metabolic transformation in below ground structures. Water transport up the xylem occurs at up to 0.8 m·s⁻¹ (Table 1) in Triticum aestivum trimmed so that only one seminal root remains^[24].

Tab.1 Speed of transmission of matter and signals in vascular plants

Process	Speed (m·s−1)	Reference
Photons in vacuo	3 × 108	[23]
Photons in plant cell	2.2 × 108	[23]
Water and solute flux root to shoot in xylem conduits	≤0.8	[24]
Water and solute flux shoot to root in phloem sieve tubes	≤1.7 × 10−3	[25]
	≤0.33 × 10−3	[26]
Basipetal polar auxin transport in parenchyma	(0.33−5) × 10−6	[27]
Solutes in cytoplasmic streaming	7 × 10−6	[28]
Pressure wave in xylem	≤1.5 × 103	[29]
Action potential in phloem	(20–50) × 10−3	[30,31]
Variation potential in phloem	(1–4) × 10−3	[32]
Ca2+ wave in vascular tissue	(1–2) × 10−3	[32]
ROS wave in vascular tissue	1 × 10−3	[32,33]

The cytosolic phloem sap in phototrophically growing vascular moves solutions from sites carrying out net photosynthetic carbon gain over the diel cycle to sites that are net chemoorganotrophic over the diel cycle, for example, growing structures above ground and, from the viewpoint of this paper, the below ground structures. There is also phloem-xylem exchange of solutes^[22]. The energization of flow from source to sink is, according to the widely accepted Münch pressure flow mechanism, the higher concentration of solutes at the source end of the pathway than at the sink end of the pathway^[34–38]. The speed (m·s⁻¹) of phloem transport is generally invariant with path length, despite limits on the maximum driving force (osmotic pressure difference), and the small observed difference in driving force with path length, between the source and sink end^[35–37] (Table 1). Decreased resistance to flow with greater path length, for example, through larger sieve pore diameter, can apparently account for the relative constancy of sap of flow with path length^[36]. If this does not always occur, it may be useful to reexamine the relay hypothesis whereby the length of sieve tubes is shorter than the path length, allowing additional energy input where solutes are unloaded at the end on one sieve tube and energized loading into the next sieve tube^[37]. The relay hypothesis only applies to the apoplasmic loading pathway. Although the most commonly recognized function of the phloem is transport of photosynthate, most commonly as sucrose, other solutes are also transported. As is discussed below, the symplasmic nature of the pathway not only constrains the amounts of certain solutes (e.g., Ca²⁺ and H⁺) that can be transported, but that there could be solutes necessarily present at relatively high concentrations (e.g., K⁺ and HPO₄^2–) because they are needed for functioning of the pathway. Importantly, relatively few plants yield pure phloem sap when cut; a cautionary case is that of the Cucurbitaceae where the copious exudate has only a small fraction of phloem sap^[39]. Excised stylets of sap-feeding insects yields pure phloem sap, but in very small quantities^[39]. Transport of solutions down the phloem occurs at up to 1.7 mm·s⁻¹ (Table 1) in T. aestivum trimmed so that only one seminal root remains^[25].

4 MATTER: CYCLING AND RECYCLING OF WATER AND NUTRIENTS BETWEEN ROOTS AND SHOOTS

Moving photosynthate and other solutes from the shoot to the roots in phloem requires movement of water as the solvent; in land plants this water must previously have been moved to the shoot in the xylem. This Münch counterflow accounts for up to 10% of the water moving up the xylem in the light, and up to 54% in the dark, in C₃ plants^[13,14,40]. The maximal 10% value for the Münch counterflow in the light is compatible with the slightly higher maximum speed of water movement in the phloem (Table 1) in view of the larger area of cross-section of xylem lumens than of sieve tubes resulting in a greater mass flow per axis in xylem than phloem^[14,17].

Cycling and recycling of elements between root and shoot plants has been reviewed^[41]. For the normal functioning of vascular land plants, C and some of the O are obtained as CO₂ from the atmosphere whereas (in decreasing order of mol of each element) H, O, N, P, S, Mg, Ca, Cl, Fe, Cu, Mn, Mo, Zn and Ni are obtained from the pedosphere. In the simplest case the soil-derived elements are either retained in the root or transported to the shoot in proportion to their requirement for growth of the two structures whereas photosynthate derived from atmospheric CO₂ and H₂O from the pedosphere is retained in the shoot or transferred to the root in proportion to the requirement for growth and maintenance of the respective structures. However, when NO₃^– is the N source and less NO₃^– is reduced and assimilated in roots than is needed for root growth, some of the organic N from NO₃^– reduced and assimilated in the shoots is cycled to roots^[42]. Also, the common S source SO₄^2– is predominantly reduced in the shoot, and organic S is cycled to the shoot^[42]. It should be that noted that excised root systems of some plants are able to assimilate both NO₃^– and SO₄^2–[43], and that some intact plants assimilate essentially all NO₃^– in their roots^[44,45].

As well as this cycling, there is also recycling of phloem-mobile ions, especially K⁺, Mg²⁺ and inorganic orthophosphate, as well as Cl^{–[41,46–51]}. In this process the ions are taken up by roots and transferred to the shoot in the xylem in excess of shoot demand, with the excess transferred back to the roots in the phloem, with some of these ions reloaded into the xylem and transported again to the shoot^[41]. Although various suggestions have been made for the role of this recycling, here the possibility is considered that the presence in, and movement along, sieve tubes of K⁺, Mg²⁺ and inorganic phosphate is a necessary consequence of the cytosolic nature of sieve tube sap whereas Cl^– and K⁺ are needed for transmission of action potentials along the sieve tube-companion cell complex.

5 MATTER: IMPLICATION FOR SOLUTE TRANSPORT OF THE CYTOSOLIC NATURE OF SIEVE TUBE SAP

The argument given here is that the cytosolic nature of the sieve tube sap means that they not only have constraints on the maximum concentration of some solutes, but also require a minimum concentration of some solutes for their functioning. All solutes in the sieve tube sap are swept along in the Münch pressure flow and consequently must be added at the source end and removed at the sink end, with exchange between the sieve tube sap and the cytosol of symplasmically connected companion cells. This statement must be modified for solutes as a whole with respect to solute exchange between sieve tube-companion cell complexes and xylem and other cells, but the concept of upper and lower limits of particular solutes is unchanged.

Dealing first with the upper limit of solute concentration in cytosol/sieve tube sap, two key solutes are H⁺ and Ca^+[52]. The free H⁺ concentration in cytosol and sieve tube sap are in the same range, between pH 7 and 8, i.e., 10–100 μmol·m⁻³, and the OH^– concentration is 100–1000 μmol·m^−3[52]. The free Ca²⁺ concentration in the plant cytosol is 50–100 μmol·m^−3[53], whereas that in the sieve tube sap, using three independent methods, is almost three orders of magnitude higher, i.e., 13–63 mmol·m^–3[54].These higher sieve tube free Ca²⁺ concentrations may have implications the functioning of the symplasmically connected companion cells^[54,55].

The capacity for transport of H⁺, OH⁻ and Ca²⁺ along phloem depends not only on the concentration of the free ions but also of the mobile buffer capacity for the ions with the over the range of free ion concentrations known within the sieve tube sap^[52]. There are no published values of buffer capacity for H⁺/OH⁻ (mol bound H⁺/OH⁻ m⁻³ (pH unit)⁻¹ or Ca²⁺ (mol bound Ca²⁺ m⁻³ (pCa unit)⁻¹). This means translating the free H⁺/OH⁻ and free Ca²⁺ concentrations relative to sucrose concentration in sieve tube sap into the capacity for H⁺/OH⁻ and Ca²⁺ transport requires assumptions as to the concentration of mobile buffers relative to that of the soluble carbohydrates providing the driving force for Münch pressure flow^[52]. Assuming no pH or pCa gradient along sieve tubes, H⁺/OH⁻ and Ca²⁺ would be loaded into the sieve tubes in parallel with the buffer compounds at the source end of the pathway, and likewise unloaded from sieve tubes at the sink end. Even considering buffering, the H⁺/OH⁻ transport capacity of phloem is insufficient to transport to the roots the excess H⁺ that would be produced from the assimilation of NH₄⁺ into organic matter (~1.3 H⁺ per N) using carbohydrate in the shoot following NH₄⁺ and Cl⁻ from the soil to the shoot in the xylem^[52,56,57]. The same is the case for the excess OH⁻ generated from the assimilation of NO₃⁻ into organic matter using carbohydrate in the shoot (~0.7 per N) following K⁺ and NO₃⁻ transport from the soil to the shoot in the xylem^[52,56,57]. These conclusions still stand when the much smaller (relative to the H⁺ or OH⁻ production in combined inorganic N assimilation) OH⁻ production during SO₄²⁻ assimilation^[56]. NH₄⁺ assimilation is limited to the root of land plants, with the excess H⁺ lost to the soil solution, with organic N needed for shoot growth transported in the xylem^[56,58]. There is no biochemical means of disposing of the quantity of H⁺ that would be generated from NH₄⁺ assimilation in the shoot^[59]. An alternative allowing mechanism allowing NH₄⁺ assimilation into organic C in the shoot, i.e., transport of one NH₄⁺ and 0.5 malate²⁻ plus ~0.3 K⁺ and 0.15 malate²⁻ up the xylem following malic acid synthesis from sucrose and CO₂ in the root, with H⁺ loss to the root medium in exchange for NH₄⁺ and K^+[56]. This does not seem to be a major pathway, although there is more malate²⁻ in the xylem sap of Ricinus communis grown with NH₄⁺ than when grown with NO₃^−[60]. Roots are the sole or predominant site of NO₃⁻ assimilation in some vascular land plants, with 0.7 OH⁻ excreted to the root medium for one NO₃⁻ and 0.3 K⁺ entering As with root NH₄⁺ assimilation, organic N used in shoot growth is transported up the xylem. In other plants shoots are the sole or predominant site of NO₃⁻ assimilation. One K⁺ and one NO₃⁻ from soil is transported up the xylem, where NO₃⁻ assimilated into organic N producing ~0.7 OH^{−[56,58,61–63]}. This is neutralized by 0.35 malate²⁻ whereas the remaining 0.3 K⁺ charge balances the net negative charge on organic compounds. The 0.7 K⁺ and 0.35 malate²⁻ is sometimes accumulated in shoot cell vacuoles^[56]. Alternatively, the 0.7 K⁺ and 0.35 malate²⁻ is transported to the root in the phloem where 0.35 malate²⁻ is metabolized generating 0.7 OH⁻ which is excreted to the root medium in exchange for 0.7 NO₃^−[56,64]. This 0.7 NO₃⁻ moves up the xylem with the 0.7 K⁺ that accompanied 0.35 malate²⁻ down then phloem, with a further 0.3 K⁺ and 0.3 NO₃⁻ from the soil solution^[56]. In the case of diazotrophic symbioses, the typical location of nodules (with the paraphyletic rhizobia or other proteobacterial symbionts such as Burkholderia) or rhizothamnia (Frankia symbionts) is below ground^[56,65]. Here the measured H⁺ excreted to the medium is greater^[62,63]; than the predicted 0.3 H⁺ per N predicted from NO₃⁻ assimilation^[56], so additional organic anion synthesis is required, some of which could be accumulated as within the plant, secreted^{[62,63,66,67]} suggested that diazotrophic nodule initiation and growth is, at least in part, regulated by phloem N. The acid-base regulation of plants with diazotrophic stem nodules (Aeschynomene, Discolobium, Neptunia and Sesbania) is not known, although the nodules can form under water as well as in humid air^[68].

The enucleate state of mature sieve tube elements in flowering plants potentially poses problems for their functioning in view of the need for proteins in sieve tubes for their functioning in the face of protein damage from reactive oxygen species (perhaps limited by hypoxia in phloem) and, in shoots, ultraviolet radiation^[55,69,70]. However, subsequent results show that some proteins (less than 20 kDa) can pass symplasmically from nucleate companion cells to enucleate sieve tube elements^[71], so it is possible that damaged proteins in sieve tube elements can be replaced. Although some mRNAs occur in sieve tube sap^[72,73], they cannot be translated in mature sieve tubes since they lack ribosomes^[73,74].

In addition to the macromolecules, small molecules are needed for the functioning of the sieve tube element-companion cell complex in maintenance of the long-distance transport pathway, loading and unloading of solutes via the apoplasmic pathway, and conversion of sucrose to raffinose and verbascose in one variant of symplasmic loading, and recouping of leaked solutes^[75]. These small molecules include K⁺ and Mg²⁺ as enzyme cofactors, and the ATP-ADP-AMP-inorganic phosphate system of energy transduction and transmission^[76,77]. It is possible that most of the functions of these small molecules occur in the nucleate companion cells, for example, apoplasmic phloem loading, recouping of leaked, sucrose, with symplasmic transfer of the sucrose to the sieve tubes. However, it seems that there is no mechanism that prevents symplasmic transfer of the catalytically active small molecules from the companion cells to the sieve tubes, and their movement is consequently by mass flow along sieve tubes. In Solanum lycopersicum the sieve tube elements and companion cells have essentially identical inside-negative electrical potential differences across their plasma membranes, i.e., are electrically connected^[78,79]. The phloem parenchyma cells have smaller electrical potential difference across the plasma membrane^[78]. Similar results were found for the difference in electrical potential difference across the plasma membrane of sieve tube elements and phloem parenchyma for R. communis and Salix alba^[80]. Electrical isolation of the sieve tube element-companion cell complex agrees with measurements of plasmodesmatal frequency distribution in the wells separating sieve tube elements and companion cells, sieve tube elements and phloem parenchyma cells, companion cells and phloem parenchyma cells, and between phloem parenchyma cells, and the spread of Lucifer yellow injected into individual cells^[75,80]. Plants with apoplasmic phloem loading have more negative electrical potentials across the sieve tube/companion cell complex plasma membrane than is the case for phloem parenchyma; in plants with symplasmic loading the electrical potential across the plasma membranes of sieve tube/companion cell complex and that of phloem parenchyma are more similar^[75].

The solutes in sieve tubes carrying solutes (and water) from shoot to root necessarily include photosynthate, usually sucrose, needed by the non-photosynthetic roots and proving much of the driving force for Münch pressure flow, and organic N and S when root-derived NO₃⁻ and SO₄²⁻ are reduced and assimilated entirely or predominantly in the shoots so that root growth requires organic N and S from the shoot^[42]. However, in addition to these compounds there are inorganic solutes derived from the root medium, for example, K⁺, Mg²⁺, inorganic phosphate and Cl^−[51]. Table 2 lists the concentration of these for inorganic ions for sieve tube sap. The flux of these solutes down the phloem appears to be a futile cycle since the roots can obtain all the K⁺, Mg²⁺, inorganic phosphate and Cl⁻ they need for their growth as well as what is transported up the xylem supplying requirements for shoot growth^[51]. In NO₃⁻-grown R. communis sieve tube sap the concentration of K⁺ is 66.0 mol·m⁻³, Mg²⁺ is 4.1 mol·m⁻³ and inorganic phosphate is 4.2 mol·m^−3[81]. Earlier work found 91.9 mol·m⁻³ K⁺, 1.5 mol·m⁻³ Mg²⁺ and 4.4 mol·m⁻³ inorganic phosphate in NH₄⁺-grown R. communis; for NO₃^–- grown R. communis the concentrations are 110 mol·m⁻³ K⁺, 1.4 mol·m⁻³ Mg²⁺ and 9.1 mol·m⁻³ inorganic phosphate^[60]. Again for NO₃⁻-grown R. communis, sieve tube sap concentrations of K⁺ is 67.1 mol·m⁻³, Mg²⁺ is 3.7 mol·m⁻³ and inorganic phosphate is 6.6 mol·m^−3[51].

Tab.2 Concentrations (mol·m−3) of K+, Mg2+, H2PO4/HPO42– and Cl– in sieve tube sap of Ricinus communis

Growth conditions	K+	Mg2+	H2PO4–/HPO42–	Cl–	Reference
External K+ is 0.4 mol·m−3 with NO3− as N source	47.0	1.5	5.0	10.9	[81]
External K+ is 1 mol·m−3 with NO3– as N source	66.0	4.1	4.2	11.4	[81]
NO3– as N source	68.1	3.9	7.6	8.9	[82]
NH4+ as N source	41.9	1.5	4.4	26.0	[60]
NO3– as N source	110.0	1.4	9.1	Not detected	[60]
NO3– as N source	67.1	3.7	6.6	12.0	[51]

In comparison, the concentration of K⁺ in the cytoplasm of a range of glycophytic flowering plants using a diversity of methods is 58–126 mol·m^{−3[83–86]}. Some of the methods, for example, compartmental analysis using ⁴²K⁺, may include the K⁺ in the other, non-vacuolar, compartments^[83]. Using ⁴²K⁺ compartmental analysis to measure cytosolic (including non-vacuolar intracellular compartments) K⁺ concentration in Hordeum vulgare roots, a range of 40–130 mol·m⁻³ was found; the lowest values correspond to high external NH₄⁺ concentrations^[87]. Using K⁺-sensitive electrodes, the activity (~90% of concentration) of K⁺ in the cytosol of H. vulgare root epidermal and cortical cells is 38–75 mol·m^–3[88]. The concentration of Mg²⁺ in the cytosol of flowering plants, determined by ³¹P nuclear magnetic resonance estimation of Mg²⁺ binding phosphate compounds, ionophores, and the kinetics of the enzyme adenylate kinase, is 0.25–0.9 mol·m^{–3[89–91]}, with rather higher free Mg²⁺ in mitochondria (1–3 mol·m⁻³) and chloroplasts (0.2−5 mol·m⁻³)^[90]. The concentration of inorganic phosphate in cytosol of Acer pseudoplatanus and A. thaliana culture cells, measured using ³¹P nuclear magnetic resonance is 0.055−0.080 mol·m^–3[92]. The most reliable estimate of cytosol Cl^– involves the use of Cl^–-specific microelectrodes yielded concentration of 11 mol·m⁻³ Cl⁻ whereas compartmental analysis of tracer Cl⁻ efflux yielded values of 5.7–21 mol·m⁻³ Cl^–; all of these values are for non-salinized glycophytes^[93,94].

Although K⁺ concentrations in cytosol and phloem are similar, Mg²⁺ and inorganic phosphate concentration in the cytosol (given above) are much lower than those in phloem (Table 2). Although the values for the sieve tube sap and the cytosol involved different plant species and (except for K⁺) different analytical methods, the differences between sieve tube sap concentration and cytosol concentrations for Mg²⁺ and, especially, inorganic phosphate suggest that there is a higher concentration of these two solutes in the sieve tube sap than would be the case if the sieve tube sap concentrations reflected that of cytosol. Although more measurements are needed, it is possible that the concentrations of inorganic phosphate and Mg²⁺ exceed in sieve tubes that needed for functioning of the symplasmically connected companion cells whereas the K⁺ concentration in sieve tube sap can be accounted for as the concentration associated with cytosol, in this case companion cell plus sieve tube element. Accordingly, the futile cycling of K⁺ down the phloem can be accounted for by the concentration needed for functioning of the sieve tube-companions cell symplasmic entity. In contrast, the greater concentration of inorganic phosphate and Mg²⁺, and, perhaps, Cl^– in sieve tube sap than in cytosol suggests that the high concentration of these solutes in sieve tube sap cannot explain futile cycling in terms of the minimum concentrations needed for companion cell-sieve tube complex function. The role of phloem in transmission of action potentials requires the presence of K⁺ and Cl^– in sieve tube sap^[30,95].

Another approach to the question of futile cycling in phloem from shoot to root is to examine the composition of sieve tube sap from phloem supplying growing vegetative and reproductive shoot structures for comparison with the composition of sieve tube sap from phloem supplying roots. Such comparisons should be within a genotype and grown under the same conditions. If there is no constraint on the composition of phloem sap, the default assumption on composition would be that the solutes in sieve tube sap moving from shoot to roots move photosynthate as sugar, and, when all NO₃^– and SO₄^2– are assimilated in shoots, organic N and S. For the phloem sap moving from transpirational termini in photosynthetic tissue to growing vegetative and reproductive structures with 24-h photosynthesis less than respiration and limited transpiration, the phloem would move the complete suite of organic C and other nutrient elements needed for growth of the structures, including the organic N needed for respiration used for growth and maintenance. However, there seem to be no such data.

A further role of K⁺, and also Cl^–, in the sieve tube sap is in action potentials transmitted along the plasma membrane of the sieve tube-companion cell complex^[30,95–98], discussed in more detail below (section 7). After initial influx of Ca²⁺ from the phloem apoplasm, the Cl⁻ efflux causes the depolarization of the inside-negative electrical potential, and subsequent K⁺ efflux causes repolarization^[30,95]. More detailed discussion of this is given below (section 7). The sieve tube-companion cell complex lack the photosynthetic apparatus that is one of the main processes in oxygenic photosynthetic organisms that require catalytic Cl^–[93,94].

Cl^–, and Na⁺, in the phloem sap have a limited role in removing NaCl that has been transported to above ground parts of glycophytes subject to salinization. Not more than 10% of the NaCl from salinized soil that reaches the shoot of glycophytes is returned to the root in the phloem^[99–102]. Osmotic and hydraulic coupling of xylem and phloem are important in salinization^[103]. There are limited data on the sieve tube sap of halophytes^[104].

6 MATTER: GAS PHASE MOVEMENT IN AERENCHYMA

Intercellular gas spaces are a key component of the functional anatomy of embryophytic land plants^[105,106]. In the context of above-below ground interactions, an important role of intercellular gas spaces is in supplying O₂ to below ground structures in waterlogged soils and flooding^[105–111]. In non-waterlogged soils the O₂ supply to below ground structures of plants is mainly by gas phase diffusion from the atmosphere through soil pores^{[105,107,112]}. The diffusion coefficient of O₂ in water is only 10⁻⁴ of that in air, and diffusive O₂ supply from the atmosphere to below ground structures through waterlogged soils at a rate sufficient for aerobic functioning of these structures is not possible^{[105,107,110]}. Plants that are tolerant of waterlogging have increased development of intercellular gas spaces in the form of aerenchyma, with individual longitudinally-running gas spaces of radii that are a significant fraction of the root radius^[107–110]. There is a negligible role of the phloem in dissolved O₂ transport from above ground to below ground plant structures^[69].

Waterlogging-tolerant plants also have barriers to lateral diffusion of O₂ from aerenchyma to waterlogged, hypoxic or anoxic, soils in the form of a suberized exodermis with lignified underlying sclerenchyma^{[108–110,113]}. This barrier presumably also limits uptake of water and nutrient solutes^[108], and may also restrict entry of toxins, such as high concentrations of Fe²⁺, generated in anoxic soil^[114]. In plants growing in non-waterlogged soil, such a barrier would limit diffusion of O₂ from the soil solution into root^[105]. O₂ leakage, and water and nutrient uptake can occur through lateral roots growing in a less deoxygenated basal region of the root and also, perhaps, through rhizodermal passage cells^[108,111].

In addition to diffusion, O₂ movement by convection (mass flow of gas) can occur when there are two or more above water structures linked by aerenchyma in a rhizome^[108,115]. Convection depends on different structure and/or function among the two (or more) above water structures, including death of one of the structures, and can be driven by differences in water vapor pressure, by thermo-osmosis, or by the venturi effect^[108,115]. Other mechanisms of mass flow can only increase the O₂ flux to below ground structures in flooded Oryza sativa by up to 6% relative to the diffusive O₂ flux^{[108,116,117]}.

As well as the role of aerenchyma in internal O₂ transport from the atmosphere (and photosynthesis in illuminated shoots) to below ground structures in waterlogging-tolerant plants, there is also a flux of CO₂ from respiration of below ground structures to above ground structures in aerenchyma. Root respiration in the presence of aerenchyma and the lateral barrier to diffusion of O₂ (and other gases) leads to a significant CO₂ flux to the shoot from root respiration^[108,111]. When the CO₂ concentration in the waterlogged soil exceeds that in the root, entry of CO₂ (not HCO₃^–) by diffusion contributes to the CO₂ flux up aerenchyma^[108,111] or in the xylem stream^[111]. Although CO₂ flux to above ground structures of the waterlogging-tolerant O. sativa from waterlogged soil in the light amounts to 20% of measured photosynthesis, leakage through lenticels mean that a much smaller fraction of the below ground derived CO₂ is assimilated in shoot photosynthesis^[111]. In emergent wetland plants CO₂ obtained through below ground parts accounts for less than 0.1% of shoot photosynthesis in the C₃ Phragmites australis^[118] and less than 0.25% in the C₃ Schoenoplectus lacuustris (as Scirpus lacustris) and C₄ Cyperus papyrus^[119]. However, up to 10% of photosynthesis by emergent leaves of the C₄ tidal marsh Sporobolus alterniflorus (as Spartina alterniflora) is supplied from below ground structures^[120]. Further work is needed. The CO₂ concentration in the xylem sap is higher than that in atmosphere-equilibrium solutions even in plants growing in non-waterlogged soils, and leakage from above ground stem structures to the atmosphere is lower in trees than in herbs^[121]. Aerenchyma is also a conduit for methane transfer to the atmosphere for plants in waterlogged soils^[108,111].

7 INFORMATION TRANSFER MECHANISMS

Apart from the well-recognized biochemical mechanisms of transfer of information between above and below ground parts of plants as hormones, peptides and small RNAs in xylem and phloem sap, and polar auxin transport from shoot to root, there are also mechanisms involving action potentials, variation potentials, Ca²⁺ waves, reactive oxygen species waves, hydrostatic waves and, possibly, light piping within the plant from the illuminated shoot to below ground parts^{[29,30,95,122–133]}.

Hydraulic pressure waves in xylem water can in principle travel at the speed of sound in water, i.e., 1500 m·s^–1[29] (Table 1). How these pressure waves in the apoplasm are perceived by target cells is not clear; mechanosensitive ion channels are a clear possibility, although knockouts of the relevant genes lacks a clear phenotype^[134]. Hydraulic transmission has been suggested as the mechanism by which, for example, sudden decreases in water potential of the root medium influences leaf growth in grasses^[122,124]. The osmoticum polyethylene glycol added to the root medium of Z. mays decreases leaf growth within 2 min of the decrease in root medium water potential^[122]. Experiments in which the decrease in xylem pressure resulting from the added osmoticant (polyethylene glycol or, more similar to nature, salt) was offset by pressurizing the root system in a pressure chamber showed that effect on leaf growth was prevented^[124]. The results were not a result of artifacts such as injection of the aqueous medium into the root gas spaces, and showed that the decreased leaf growth was related to the decreased pressure in the xylem rather than a chemical messenger^[124]. However, these experiments do not appear to exclude the possibility of a pressure difference-induced electrical signal transmitted along the phloem. Also, a sudden increase in root medium osmotic potential (decreased water potential) of the magnitude used in these experiments do not occur in the natural environment^[122,124]. Whether acoustic signals, of the kind used by investigators to detect embolism of individual xylem conduits, are used for within-plant signaling deserves investigation^[135].

Light travels at 2.2 × 10⁸ m·s⁻¹ in plant cells^[23] (Table 1). Light piping through files of cells is known to occur in vascular land plants^[16,17,136]. Light can also penetrate to a limited extent into soil^[137]. For both plants and soil exposed to sunlight, far-red light penetrates further than red light, which penetrates further than blue light^[137]. Correspondingly, the blue light absorbing photoreceptor, phototropin, is expressed in roots near the hypocotyl whereas the red to far-red absorbing phytochromes are expressed near the root apex^[130,137]. Phytochrome in root tips was activated by light incident on shoots and transmitted along the root tip, with photomorphogenetic effects on growth and gravitropism; however, the length of the roots is not stated, so the distance in the root over which light sufficient to act on phytochrome is transmitted is not known^[130]. A retinal binding protein is involved in the oscillatory mechanism (root clock) that regulates the origin of lateral roots in A. thaliana^[138], but there is no evidence as to a role of light absorption by retinal such as occurs in phototaxis by some flagellate algae^[139].

Other means of information transfer are the Ca²⁺ wave and the reactive oxygen species (ROS) wave^{[126–128,131,132,140]}. The Ca²⁺ wave involves a two-pore Ca²⁺ channel in the plasma membrane, and also glutamate receptor-like proteins 3.3 and 3.6, and travels at about 0.4 mm·s^{−1[127,128,141]} (Table 1). The ROS wave involves increased activity of a NADPH oxidase generating ROS in the apoplasm, and travels at about 1.4 mm·s^{−1[126,128,140]} (Table 1). These waves may involve electrical transmission resulting from plasma membrane depolarization by apoplasmic ROS or by Ca^{2+[126,131,142]}.

The Ca²⁺ wave and the ROS wave are transmitted faster than the mass flow in most measurements in the phloem (0.33 mm·s^−1[26,35]: but at a similar speed to the highest measured value of 1.7 mm·s^−1[25] (Table 1)). Also, they can move in either direction rather than just shoot to root as for mass flow in the phloem. The Ca²⁺ and ROS waves move much more rapidly than the 7.2 μm·s⁻¹ speed of cytoplasmic streaming, in A. thaliana^[28] (Table 1). Even when A. thaliana myosin XI-2 was made chimeric with the giant-celled alga, Chara corallina, myosin XI, giving streaming speeds in C. corallina up to 100 μm·s⁻¹, the streaming speed of A. thaliana is only increased to 16 μm·s^−1[28]. Finally, diffusion^[143,144] of Ca²⁺ or ROS to activate, respectively, Ca²⁺ channels and NADPH oxidases, increasing the concentration of Ca²⁺ in the cytosol and of ROS in the apoplasm, in propagating, the wave by a relay mechanism, is far to too slow to account for the wave speeds.

Electrical transmission of information can occur as action or variation potentials along the phloem, involving depolarization of the inside-negative electrical potential difference by limited Ca²⁺ influx and greater Cl⁻ efflux, followed by repolarization K⁺ efflux^{[30,95–98,123]}. Table 2 shows that, at least in R. communis, the K⁺ concentration is sieve tube sap ranges from 47 to 110 mol·m⁻³ whereas the Cl⁻ concentration is 8.9−26 mol·m⁻³ with the exception of the finding^[60] for NO₃⁻-grown R. communis of no detectable Cl⁻. Much work has been carried out on action potentials using giant intermodal cells of the Characeae; these are algal members of the Streptophyta, the clade to which flowering plants belong^[144,145]. However, internally perfused internodal cells of Chara and showed that normal action potentials were found with cytosol Cl⁻ concentrations from 0.01 to 29 mol·m⁻³ Cl^−[146]. If a cytosol Cl⁻ concentration as low as 0.01 mol·m⁻³ allows action potentials in R. communis, then the absence of detectable Cl⁻ in the sieve tube sap NO₃⁻ grown R. communis^[60] may have been compatible with the occurrence of 0.01 mol·m⁻³ Cl⁻ in the sieve tube sap. More work is needed.

The speed of transmission of action potentials in phloem is 20−50 mm·s^−1[30,98] (Table 1) whereas angiosperm tree and herb phloem mass flow is at 0.17 mm·s^−1[26,35] and up to 1.7 mm·s^−1[25] (Table 1). Variation potentials travel at 1−4 mm·s^−1[32] (Table 1). Although the direction of mass flow in phloem is determined by source-sink relations, action potentials can be transmitted in either direction. However, action potentials carry little information relative to the multitude of informational molecules transported by mass flow in phloem and, to a lesser extent, in xylem.

Importantly, Fichman and Littler^[132] show that glutamate receptor-like proteins 3.3 and 3.6 integrate electrical, Ca²⁺, reactive oxygen species and hydraulic systemic signals. Also, reactive oxygen species-enhanced signaling involves a plasmodesma-located protein, providing an additional link between systemic signaling and symplasmic transport^[147].

Transport of low molecular mass hormones, peptides and miRNA can occur in xylem and/or phloem^{[132,148–150]}; there is also the possibility that the increase in root xylem sap pH in response to drought also has a signaling function^[151]. Auxin (indoleacetic acid) is subject to shoot-root (basipetal) polar transport in parenchyma at 0.33−5 μm·s^−1[27] (Table 1). The polar transport involves indoleacetate⁻:H⁺ cotransport into cells at the source end, and indoleacetate⁻ efflux through PIN channels driven by the inside-negative electrical potential difference across the plamalemma generated by H⁺ efflux using a H⁺ P-ATPase. The higher speeds may require cytoplasmic streaming, though inhibition by streaming inhibitors such as cytochalasin is complicated by effects on the polar distribution of transport proteins in the upstream and downstream ends of the cell^[152].

Long-distance transport of hormones in xylem and/or phloem has been reported for abscisic acid (ABA), cytokinin, strigolactone, and the precursor, 1-aminocyclopropane-1-carboxylic acid (ACC) of the gaseous ethylene^[149]. Although the ABA transported to shoots in the xylem in response to drought has been reported as being synthesized in roots, movement of ABA up the xylem to stomata would give slow stomatal response to changes in the root medium^[153], and later results show that ABA can be synthesized in shoot vascular tissue in response to drought, and ABA can be synthesized in guard cells in response to low relative humidity^[149]. Different cytokinins can transported acropetally in the xylem and basipetally in the phloem^[149]. ACC can also move in both xylem and phloem^[149].

Variation in xylem sap pH in response to below ground conditions might transmit information to the shoot, either directly or by influencing the uptake in the shoot of hormones, especial those that are weak acids and bases such as ABA and cytokinins^{[151,154–156]}. Drought increases the xylem sap pH which would, if reflected in the leaf apoplasm, increase stomatal aperture, but, via effects on ABA distribution between apoplasm and symplasm in the leaf leading to a higher apoplasmic ABA concentration, and thereby causes stomatal closure^[155]. Increased xylem pH is apparently also involved in decreased stomatal aperture when plants are grown with conspecifics and competition for water and nitrogen are eliminated^[157].

Finally, innovative use in information transfer has been made of the flowering plant holoparasite, Cuscuta campestris, that parasitizes a range of euphyllophytes through haustoria on above ground parts^[97,158]. For a C. campestris individual parasitizing two hosts of different species it was) shown that systemic wound signals could be transmitted between dicot and monocot hosts, and between dicot and fern hosts, consistent with similar signals being used throughout euphyllophytes (ferns and seed plants)^[158]. A systemic signal of N-deprivation in Glycine max was transmitted via C. campestris to N-replete Cucumis sativa where NO₃⁻ uptake was stimulated, again showing commonality of the N-deprivation signal of the two hosts^[97].

8 ECOSYSTEM SERVICES

Ecosystem services covers a wide range of ecosystem effects that influence human beings^[159]. Here agroecosystems are emphasized, where obvious benefits to humanity are the food, fiber and fuel that have obvious economic value, and clearly involve above and below ground interactions irrespective of whether the harvested crop is above ground, such as T. aestivum grain, or below ground, such as Solanum tuberosum tubers. However, the other outcomes of agroecosystems that influence humanity need be considered.

Organic C from photosynthesis transferred to below ground structures, and subsequently respired to CO₂ can contribute to CO₂ sequestration as inorganic carbon in soil via recalcitrant plant components, and as HCO₃⁻ by respired CO₂ whose diffusive loss to the atmosphere is limited, chemically weathering Ca and Mg silicates and carbonates in Eq. (1) (calcite), Eq. (2) (forsterite) and Eq. (3) (anorthite)^[160].

(1)$\mathrm{C}\mathrm{a}\mathrm{C}{\mathrm{O}}_3+\mathrm{C}{\mathrm{O}}_2+{\mathrm{H}}_2\mathrm{O}\text{→}\mathrm{C}{\mathrm{a}}^{2+}+2\mathrm{H}\mathrm{C}{\mathrm{O}}_3^{-}$

(2)$\mathrm{M}{\mathrm{g}}_2\mathrm{S}\mathrm{i}{\mathrm{O}}_4+4\mathrm{C}{\mathrm{O}}_2+4{\mathrm{H}}_2\mathrm{O}\text{→}2\mathrm{M}{\mathrm{g}}^{2+}+4\mathrm{H}\mathrm{C}{\mathrm{O}}_3^{-}+{\mathrm{H}}_4\mathrm{S}\mathrm{i}{\mathrm{O}}_4$

(3)$\mathrm{C}\mathrm{a}\mathrm{A}{\mathrm{l}}_2\mathrm{S}{\mathrm{i}}_2{\mathrm{O}}_8+2\mathrm{C}{\mathrm{O}}_2+3{\mathrm{H}}_2\mathrm{O}\text{→}\mathrm{C}{\mathrm{a}}^{2+}+2\mathrm{H}\mathrm{C}{\mathrm{O}}_3^{-}+\mathrm{A}{\mathrm{l}}_2\mathrm{S}{\mathrm{i}}_2{\mathrm{O}}_5(\mathrm{O}\mathrm{H}{)}_4$

Such weathering was greatly enhanced with the origin of rooted embryophytic plants over 400 million years ago^[161], and is a major source of HCO₃⁻ to the ocean. However, this oceanic HCO₃⁻ is not a very long-term CO₂ sink since biological CaCO₃ precipitation occurs with release of CO₂ (Eq. (4)), the reverse of Eq. (1):

(4)$\mathrm{C}{\mathrm{a}}^{2+}+2\mathrm{H}\mathrm{C}{\mathrm{O}}_3^{-}\text{→}\mathrm{C}\mathrm{a}\mathrm{C}{\mathrm{O}}_3+\mathrm{C}{\mathrm{O}}_2+{\mathrm{H}}_2\mathrm{O}$

However, enhanced HCO₃⁻ production is soil water by addition of silicate minerals to soils is a plausible role of below ground metabolism of plants in CO₂ sequestration in the ocean, unless the rate of calcification, largely by coccolithophores and foraminifera, in the ocean increases to an extent that compensates for HCO₃⁻ input^{[160,162,163]}.

Storage of CO₂ as organic C in soil can occur as recalcitrant polymers from dead plants such as lignin and, from above ground parts, cutin. Polymeric carbohydrates (pectins, arabinogalactans, xyloglucans from plant roots, and the glomalin thought to be produced by root-symbiotic arbuscular mycorrhizal fungi^[164] are important agents of soil aggregation^[165]. Production of combined N by symbiotic and non-symbiotic diazotrophy, and mobilization and movement through soil of combined N, P and Fe, in the soil, all depend on organic C from shoot photosynthesis^[166,167]. All of these effects are larger per m² soil area with vascular plant vegetation, with their greater volumetric penetration into the soil, than with cyanobacteria, algae and bryophytes^{[161,165,168]}. To the extent that crops have a higher productivity than the vegetation that would otherwise occur on that area there is the potential for enhanced soil ecosystem services; however, fertilizer input can alter the ecosystem processes. It is not clear what effect the soil aggregates ultimately produced by plant organic C have on water flow in soil and allocation of precipitation into liquid water transfer to ground water, streams and the ocean, to evaporation, and to transpiration, with their effects on humanity through the supply of potable water and on local weather^[169,170].

One important ecosystem service related to organic C transport from shoots to below ground parts is accumulation of organic C in soil^{[19,171–175]}. The decrease in atmospheric CO₂ and CH₄ in the pre-agricultural Holocene was reversed about 8000 years ago for CO₂, paralleling the widespread occurrence of agriculture and associated deforestation, and for CH₄ about 5000 years ago with introduction of irrigation rice production and domesticated^[176]. Agriculture, and wetland degradation, have been causally related to these reversals of decreases in atmospheric greenhouse gas^[171]. The decrease in organic C in soil and above ground biomass over the past 8000 years is 38 Pmol C^[171]. Agriculture has resulted in a global reduction of 9.7 Pmol organic C in the top 2 m of global soils, i.e., an ecosystem disservice^[172]. The global reduction of soil organic C from a meta-analysis of peer-reviewed literature yielded a value of 11.3 Pmol C^[173]. Methods have been suggested by which this loss of soil organic C in agricultural land could be reversed^[173,174]; however, the warming resulting from additional CO₂ and CH₄ from agriculture has exerted a positive feedback by decreased CO₂ solubility in a warmer ocean^[175].

The discussion of the impact of agriculture on sequestration in soil of organic C produced from atmospheric CO₂ above shows that soil organic C is significantly below what would have occurred from continuation of the pre-agricultural vegetation.

9 CONCLUSIONS

Some aspects of the balance of above and below ground growth of vascular land plants can be rationalized by the models considering the rate of acquisition of resources supplied from the aerial environment (light and CO₂) and the soil (H₂O, essential elements other than C and some O). However, such models do not account, for example, for storage in reproductive structures in shoots and below ground storage structures, and information flow as well as resource flow between above and below ground plant parts is needed. There is wide acceptance of the cohesion-tension hypothesis for the mechanism of transpiratory flow of water and dissolved solutes derived from the soil from roots to shoot in the xylem, and of the Münch pressure flow of transport of dissolved carbohydrates and other solute from shoot to below ground structures in the phloem. However, some aspects need further research, such as the possibility of active water transport in contributing to root pressure under low transpiration conditions. The cytosolic nature of the contents of sieve tube elements and symplasmically connected companion cells imposes constraints on the transport from shoots to roots of buffered H⁺/OH⁻ generated by metabolism in the shoot with implications for the location of the assimilation of inorganic N. A further, less investigated, constraint on phloem transport is that there is a minimum concentration of cytosolic ions such as K⁺, Mg²⁺ and H₂PO₄⁻/HPO₄²⁻ as well as ATP and ADP to operate and maintain the transport system, and mass flow of solution means that these solutes must originate in the shoot and be delivered to the roots. This explains, in part, the futile cycling of root-derived nutrients returned from the shoot to the root, and constraints on how much NaCl delivered to the shoot in the xylem from saline soils can be returned to the roots. Aerenchyma, and barriers to radial O₂ loss, are important in supplying O₂ to below ground structures in waterlogged soil.

Above-below ground interactions also involve information transfer used for integration of growth of plant parts and, in the short-term, communication of abiotic and biotic damage and environmental changes. As well as the obvious transfer of phytohormones in xylem and phloem, faster communication can occur through pressure waves and electrical (variation and action potentials) signals, the latter apparently including Ca²⁺ waves and ROS waves.

Agriculture has decreased the ecosystem benefit of organic C accumulation in soil and consequently decreased CO₂ removal from the atmosphere, i.e, an ecosystem disservice The incorporation of atmospheric CO₂ into soil, and eventually into the ocean, can be stimulated by spreading particulate silicate minerals on agricultural soils. This accumulation of atmospheric C as dissolved inorganic C is an ecosystem service.

References

[1] WeigeltA, JolliffeP. Indices of plant competition. Journal of Ecology, 2003, 91( 5): 707– 720

[2] BedoussacL, JustesE. A comparison of commonly used indices for evaluating species interactions and intercrop efficiency: application to durum wheat–winter pea intercrops. Field Crops Research, 2011, 124( 1): 25– 36

[3] Makowski D, Piraux F, Brun F. From experimental network to meta-analysis-methods and applications with R for agronomic and environmental sciences. Springer, 2019

[4] ConnollyJ, GomabH C, RahimK. The information content of indicators in intercropping research. Agriculture, Ecosystems & Environment, 2001, 87( 2): 191– 207

[5] Li L, Zhang L, Zhang F. Crop mixtures and the mechanisms of overyielding. In: Levin S A, ed. Encyclopedia of Biodiversity (2nd ed.). Waltham: Academic Press, 2013, 382–395

[6] TilmanD. Benefits of intensive agricultural intercropping. Nature Plants, 2020, 6( 6): 604– 605

[7] MalézieuxE, CrozatY, DuprazC, LauransM, MakowskiD, Ozier-LafontaineH, RapidelB, deTourdonnet S, Valantin-MorisonM. Mixing plant species in cropping systems: concepts, tools and models. A review. Agronomy for Sustainable Development, 2009, 29( 1): 43– 62

[8] YuY, Stomph T J, MakowskiD, van der WerfW. Temporal niche differentiation increases the land equivalent ratio of annual intercrops: A meta-analysis. Field Crops Research, 2015, 184 : 133– 144

[9] Martin-GuayM O, PaquetteA, DuprasJ, RivestD. The new Green Revolution: Sustainable intensification of agriculture by intercropping. Science of the Total Environment, 2018, 615 : 767– 772

[10] XuZ, Li C, ZhangC, YuY, van der Werf W, ZhangF. Intercropping maize and soybean increases efficiency of land and fertilizer nitrogen use; A meta-analysis. Field Crops Research, 2020, 246 : 107661

[11] FengC, SunZ X, ZhangL, FengL S, ZhengJ M, BaiW, Gu C F, WangQ, XuZ, van der Werf W. Maize/peanut intercropping increases land productivity: a meta-analysis. Field Crops Research, 2021, 270 : 108208

[12] LiC, Hoffland E, KuyperT W, YuY, Zhang C, LiH, ZhangF, van der WerfW. Syndromes of production in intercropping impact yield gains. Nature Plants, 2020, 6( 6): 653– 660

[13] MeadR, WilleyR W. The concept of a ‘Land Equivalent Ratio’ and advantages in yields from intercropping. Experimental Agriculture, 1980, 16( 3): 217– 228

[14] WangQ, BaiW, Sun Z, ZhangD, ZhangY, WangR, EversJ B, StomphT J, van der WerfW, FengC, ZhangL. Does reduced intraspecific competition of the dominant species in intercrops allow for a higher population density?. Food and Energy Security, 2021, 10( 2): 285– 298

[15] SpittersC J T. An alternative approach to the analysis of mixed cropping experiments. 1. Estimation of competition effects. Netherlands Journal of Agricultural Science, 1983, 31( 1): 1– 11

[16] LoreauM, HectorA. Partitioning selection and complementarity in biodiversity experiments. Nature, 2001, 412( 6842): 72– 76

[17] LiC, Hoffland E, KuyperT W, YuY, Li H, ZhangC, ZhangF, van der WerfW. Yield gain, complementarity and competitive dominance in intercropping in China: A meta-analysis of drivers of yield gain using additive partitioning. European Journal of Agronomy, 2020, 113 : 125987

[18] ZhangL, van der WerfW, ZhangS, LiB, Spiertz J H J. Growth, yield and quality of wheat and cotton in relay strip intercropping systems. Field Crops Research, 2007, 103( 3): 178– 188

[19] van OortP A J, GouF, Stomph T J, van der WerfW. Effects of strip width on yields in relay-strip intercropping: A simulation study. European Journal of Agronomy, 2020, 112 : 125936

[20] WangR, SunZ, Zhang L, YangN, FengL, BaiW, Zhang D, WangQ, EversJ B, LiuY, Ren J, ZhangY, van der WerfW. Border-row proportion determines strength of interspecific interactions and crop yields in maize/peanut strip intercropping. Field Crops Research, 2020, 253 : 107819

[21] Harper J L, Ltd H P. Population biology of plants. London: Academic Press, 1977

[22] WeinerJ, FreckletonR P. Constant final yield. Annual Review of Ecology, Evolution, and Systematics, 2010, 41( 1): 173– 192

[23] Bastiaans L, Storkey J. Descriptive and mechanistic models of crop–weed competition. In: Hatcher P E, Froud-Williams R J, eds. Weed Research: Expanding Horizons, 2017, 33–60

[24] Trenbath B R. Biomass productivity of mixtures. In: Brady N C, ed. Advances in Agronomy. Academic Press, 1974, 26: 177–210

[25] Yu Y. Crop yields in intercropping: meta-analysis and virtual plant modelling. Dissertation for the Doctoral Degree. The Netherlands: Wageningen University, 2016

[26] CardinaleB J, WrightJ P, CadotteM W, CarrollI T, HectorA, SrivastavaD S, LoreauM, WeisJ J. Impacts of plant diversity on biomass production increase through time because of species complementarity. Proceedings of the National Academy of Sciences of the United States of America, 2007, 104( 46): 18123– 18128

[27] BerghuijsH N C, WangZ, StomphT J, WeihM, Van der WerfW, VicoG. Identification of species traits enhancing yield in wheat-faba bean intercropping: development and sensitivity analysis of a minimalist mixture model. Plant and Soil, 2020, 455( 1−2): 203– 226

[28] de WitC T. Resource use efficiency in agriculture. Agricultural Systems, 1992, 40( 1-3): 125– 151

[29] MonteithJ L, MossC J, CookeG W, PirieN W, BellG D H. Climate and the efficiency of crop production in Britain. Philosophical Transactions of the Royal Society of London. Series B: Biological Sciences, 1977, 281( 980): 277– 294

[30] van IttersumM K, RabbingeR. Concepts in production ecology for analysis and quantification of agricultural input-output combinations. Field Crops Research, 1997, 52( 3): 197– 208

[31] GouF, van Ittersum M K, SimonE, LeffelaarP A, van der PuttenP E L, ZhangL, van der WerfW. Intercropping wheat and maize increases total radiation interception and wheat RUE but lowers maize RUE. European Journal of Agronomy, 2017, 84 : 125– 139

[32] ZhuJ, van der Werf W, AntenN P R, VosJ, Evers J B. The contribution of phenotypic plasticity to complementary light capture in plant mixtures. New Phytologist, 2015, 207( 4): 1213– 1222

[33] GouF, van Ittersum M K, Couëdel A, ZhangY, WangY, vander Putten P E L, ZhangL, vander Werf W. Intercropping with wheat lowers nutrient uptake and biomass accumulation of maize, but increases photosynthetic rate of the ear leaf. AoB Plants, 2018, 10( 1): ply010

[34] MaoL, Zhang L, LiW, van der WerfW, SunJ, Spiertz H, LiL. Yield advantage and water saving in maize/pea intercrop. Field Crops Research, 2012, 138 : 11– 20

[35] MaL, Li Y, WuP, ZhaoX, ChenX, GaoX. Coupling evapotranspiration partitioning with water migration to identify the water consumption characteristics of wheat and maize in an intercropping system. Agricultural and Forest Meteorology, 2020, 290 : 108034

[36] TanM, Gou F, StomphT J, WangJ, YinW, Zhang L, ChaiQ, van der WerfW. Dynamic process-based modelling of crop growth and competitive water extraction in relay strip intercropping: model development and application to wheat-maize intercropping. Field Crops Research, 2020, 246 : 107613

[37] MorrisR A, GarrityD P. Resource capture and utilization in intercropping: water. Field Crops Research, 1993, 34( 3−4): 303– 317

[38] RenJ, Zhang L, DuanY, ZhangJ, EversJ B, ZhangY, SuZ, van der Werf W. Intercropping potato ( Solanum tuberosum L.) with hairy vetch ( Vicia villosa) increases water use efficiency in dry conditions. Field Crops Research, 2019, 240 : 168– 176

[39] TangX, ZhangC, YuY, Shen J, van der WerfW, ZhangF. Intercropping legumes and cereals increases phosphorus use efficiency; a meta-analysis. Plant and Soil, 2021, 460( 1−2): 89– 104

[40] GouF, van Ittersum M K, WangG, van der PuttenP E L, van der WerfW. Yield and yield components of wheat and maize in wheat–maize intercropping in the Netherlands. European Journal of Agronomy, 2016, 76 : 17– 27

Compliance with ethics guidelines

Wopke van der Werf, Lizhen Zhang, Chunjie Li, Ping Chen, Chen Feng, Zhan Xu, Chaochun Zhang, Chunfeng Gu, Lammert Bastiaans, David Makowski, and TjeerdJan Stomph

RIGHTS & PERMISSIONS

Frontiers of Agricultural Science and Engineering