Disposable electrochemical flow cells for catalytic adsorptive stripping voltammetry (CAdSV) at a bismuth film electrode (BiFE)

Catalytic adsorptive stripping voltammetry (CAdSV) has been demonstrated at a bismuth film electrode (BiFE) in an injection-moulded electrochemical micro-flow cell. The polystyrene three-electrode flow cell was fabricated with electrodes moulded from a conducting grade of polystyrene containing 40% carbon fibre, one of which was precoated with Ag to enable its use as an on-chip Ag/AgCl reference electrode. CAdSV of Co(II) and Ni(II) in the presence of dimethylglyoxime (DMG) with nitrite employed as the catalyst was performed in order to assess the performance of the flow cell with an in-line plated BiFE. The injection-moulded electrodes were found to be suitable substrates for the formation of BiFEs. Key parameters such as the plating solution matrix, plating flow rate, analysis flow rate, solution composition and square-wave parameters have been characterised and optimal conditions selected for successful and rapid analysis of Co(II) and Ni(II) at the ppb level. The analytical response was linear over the range 1 to 20 ppb and deoxygenation of the sample solution was not required. The successful coupling of a microfluidic flow cell with a BiFE, thereby forming a “mercury-free” AdSV flow analysis sensor, shows promise for industrial and in-the-field applications where inexpensive, compact, and robust instrumentation capable of low-volume analysis is required.
Introduction
Mercury electrodes, normally in the form of mercury films for flow systems, have been widely used for performing stripping voltammetry as a result of their high sensitivity and reproducibility, but, due to increasing concern and legislation regarding the toxicity of mercury, alternative working electrode (WE) materials have been investigated. Metals such as Au [1, 2], Ag [2, 3], Ir [4] and even W [5] as well as bare and coated carbon electrodes [6, 7] have all been investigated for stripping analysis of metals.
In 2000, bismuth-film electrodes [8] were mooted as a replacement for mercury-film electrodes, as bismuth exhibits negligible toxicity and is more environmentally friendly in comparison. Mercury use is increasingly regulated, motivating the search for mercury-free methods. For example, Sweden banned the sale of switches, apparatuses and instruments containing mercury in 1993 (Ordinance SFS 1991:1290) and instructed the Swedish Chemicals Inspectorate (KemI) to look into a total ban on the handling of mercury in Sweden, including chemicals used for analysis. The Inspectorate proposed that mercury for chemical analysis should be exempted from the general ban until 31st December 2008 in order to enable the development and implementation of mercury-free analysis methods [9]. In 2005 the EU commission proposed a ban on mercury exports by 2011 as part of a strategy to reduce mercury emissions and protect against exposure [10], and recently MEPs voted to bring this date forward to 2010 [11].
A variety of substrates have been used for bismuth films. While platinum [12–14] and gold [13] have both been investigated, the majority of films have been deposited on carbon substrates. Most studies have used glassy carbon [8, 13, 15–32], though wax-impregnated graphite [22, 24], pencil-lead graphite [33], carbon paste [13, 34–36], screen-printed carbon inks [21, 37, 38], carbon fibres [8, 13, 14] and boron-doped diamond [39] have all been employed as substrates. The use of Bi2O3 containing pastes [34, 40] and a bismuth bulk electrode [41] have also been reported.
The use of BiFE has recently been reviewed [42]. Bismuth-film electrodes have mainly been applied to anodic stripping voltammetry (ASV) and adsorptive stripping voltammetry (AdSV) techniques for metal ion analysis, although cathodic detection of nitrophenols [17, 27, 41] and amperometric detection of glucose [12] have also been reported. Cd [28], Pb [18, 37], Zn [31, 32] and mixtures of all three ions [8, 20, 22, 23, 33, 36, 41] have been the most extensively studied by ASV. Tl, Cu [16], In [16, 43], Mn [39] and Sn [44] have also been analysed. Fewer reports have been made employing AdSV, though Ni [15] and Co [28–30] and Ni with Co [24, 26] have been examined using dimethylglyoxime (DMG). Cr [21, 25, 45], U [46, 47], Al [48], V [49] and Mo [50] analysis by AdSV on bismuth-film electrodes has also been performed.
Stirred solutions have normally been used, though Economou et al. have reported the use of a rotating disc electrode [22–25] and a Nafion-coated BiFE in a thin layer flow cell (0.5 mm spacer) for sequential injection-ASV of metals [51, 52]. Hutton et al. generated an in-situ BiFE in a thin layer (0.3 mm spacer) flow cell (Bioanalytical Systems Inc., West Lafayette, IN, USA) for flow injection analysis of nitrophenols with cathodic amperometric detection [27].
Adsorptive stripping voltammetry of DMG-complexed Co and Ni has found widespread use in trace metal analysis. The enhancement of the Co-DMG response using nitrite, i.e. catalytic adsorptive stripping voltammetry (CAdSV), has also been the subject of several publications, for example van den Berg et al. [53, 54] and the review of catalytic systems by Bobrowski and Zarębski [55]. CAdSV of Co on a BiFE was reported by Krolicka et al. [29, 30] using an ex situ plated film on glassy carbon. Most BiFEs have been plated from acetic buffers, although Krolicka et al. used a plating solution of Bi(NO3)3 (0.02 M), LiBr (0.5 M), and HCl (1 M) with a plating potential of −0.25 V for 45 s [addition of bromide and strong acid enabled a highly concentrated Bi(III) plating solution to be produced]. Stripping was typically carried out in 0.1 M ammonia buffer (pH 9.2) containing DMG (1 × 10−4 M) and NaNO2 (0.5 M). Voltammograms were recorded in the differential pulse mode and a 15-fold enhancement of the Co-DMG stripping signal was reported.
Recently the authors reported the use of 40% carbon fibre-filled nylon 6/6 injection-moulded electrodes as working electrodes for the determination of Cu(II) by CV and ASV techniques [56].
In this work, the electrodes were moulded from a 40% carbon fibre-filled polystyrene, which was also found to be suitable for performing electrochemical detection. This paper presents the first report of the use of CAdSV at an in-line plated BiFE in an integrated three-electrode injection-moulded plastic micro flow cell.
Experimental
Instrumentation
Voltammetric measurements were performed using an Autolab electrochemical system (PSTAT 10, Eco Chemie B.V., Utrecht, The Netherlands) controlled with GPES software (v4.6, Eco Chemie) running on a PC.
Rotating disc electrode experiments
The rotating disc electrode (RDE) was a glassy carbon electrode (6.1204.000, Metrohm, Buckingham, UK) 3 mm in diameter controlled with a variable speed (500–3000 rpm) electrode rotator (Part No. 628-10, Metrohm). The disc electrode was polished with an alumina slurry (0.3 μm Al2O3, Metrohm) before use. A standard (12 mm in diameter) glass double junction Ag/AgCl/KCl (3M) reference electrode was used (CR4/DJ/AG, Thermo Electron, Auchtermuchty, Fife, UK) with a Pt wire auxiliary 0.5 mm in diameter (99.9%, Aldrich, Gillingham, UK). The electrodes were mounted in a small-volume 50-ml beaker used as an electrochemical cell.
Flow cell experiments
A gravity-fed flow system was used comprising a reservoir formed from a disposable syringe body (20 ml), and narrow-bore PTFE tubing (0.032 in. i.d., Lee Products, Gerrards Cross, UK). The flow was controlled with a manual two-way valve (Part No. 2420, Omnifit, Cambridge, UK). Barbed (1/16″) polypropylene female Luer fittings (Cole-Parmer, Hanwell, London, UK) enabled connection of the flow system to the injection-moulded flow cell. Electrical connection to the Autolab electrochemical system was made using crocodile clips.
Chip fabrication
The injection-moulded flow cells were produced in-house. Figure 1 depicts the fabricated device showing the three integrated electrodes (WE, reference and auxiliary). Solid models created using AutoCAD (Mechanical Desktop 2004, Autodesk, San Jose, CA, USA) were converted into macro commands for a CNC milling machine (Datron CAT3D M6, Datron Technology, Milton Keynes, UK) to produce the milled aluminium moulds for the injection-moulding machines (Babyplast 6/6 or 6/10, Cronoplast SL, Barcelona, Spain). A four-stage process was required to manufacture a ready-to-use device. The polymer electrodes were injection moulded (moulding temperature of 220 °C) from a conducting polymer (40% carbon-fibre-filled high-impact polystyrene (HIPS), RTP 487, RTP Company (UK) Plastics Ltd., Bury, UK). The reference electrodes were coated with Ag (Ag evaporation slug, 99.99%, Aldrich) using an e-beam evaporator (Auto500 with EB3 e-beam unit, Edwards High Vacuum International, Crawley, West Sussex, UK) to a thickness of 125 nm. The WE, reference and auxiliary electrodes were then incorporated into the flow cell by an overmoulding procedure. The mould cavity for the flow cell design comprises a base plate into which the preformed electrodes are inserted in the appropriate recesses and a top plate with a relief pattern that forms the flow channel and integrated fluidic connectors. The flow cell was moulded (moulding temperature of 220 °C) from crystal polystyrene (Northern Industrial Plastics Ltd. Chadderton, UK). Using a 500-W ultrasonicwelder (Mini delta 9500, FFR Ultrasonics Ltd., Queniborough, Leicestershire, UK), with a weld time of 1 s and a hold time of 2 s, the top and base plates were sealed together to form a complete and ready-to-use device. The flow channel dimensions were 1 mm wide, 1 mm deep and 30 mm long. The working and auxiliary electrodes were 5 mm wide, the reference was 2 mm wide and the spacing between them was 2 mm.
Fig. 1The injection-moulded electrochemical micro flow cell. Here (A) is the Ag-coated electrode, which forms the reference (Ag/AgCl) element; (B) is the working electrode, on which the bismuth film electrode is formed; (C) is the auxiliary electrode; (D) is the device inlet and (E) is the device outlet. The flow channel is 1 mm × 1 mm and 30 mm long
Reagents and solutions
All aqueous solutions were prepared using >18MΩ water (Elga Maxima Ultra Pure, Vivendi, High Wycombe, UK). The acetate buffer was prepared from acetic acid (glacial 99.8% AnalaR grade, BDH, Poole, UK) and sodium acetate (99+% ACS grade, Aldrich, Gillingham, UK). The ammonium buffer was prepared from ammonium chloride (99.8% AnalaR grade, BDH) and ammonium hydroxide (28–30% NH3 ACS grade, Aldrich). Bismuth plating solutions were prepared with bismuth(III) nitrate pentahydrate (99.999%, Aldrich,) either in acetate buffer or with lithium bromide (99+%, Aldrich) in hydrochloric acid (1 M volumetric standard, Riedel-de Haën, Gillingham, UK), as described in the “Procedure” section. The mercury plating solution was prepared in nitric acid from a 1000 ppm mercury(II) chloride (98+%, Aldrich) stock solution, as described in the “Procedure” section. Sample solutions were made from cobalt (995 μg/l, Aldrich) and nickel (1 g/l, Fluka, Gillingham, UK) atomic absorption standards with dimethylglyoxime (99+%, Acros, Loughborough, UK) and sodium nitrite (97+%, Aldrich). The saturated AgCl solution was prepared from silver nitrate (99.9% AnalaR grade, BDH) in potassium chloride (99.5% AnalaR grade, BDH). Nitric acid (2 M volumetric standard, Riedel-de Haën) was used for electrode cleaning, as described in the “Procedure” section.
Procedure
Formation of the Ag/AgCl miniaturised reference electrode
Using an external auxiliary platinum wire located in the inlet of the microdevice, the Ag electrode was coated in AgCl (in saturated KCl solution that was also saturated with AgCl through the addition of a small quantity of AgNO3) at 2 V for 3 s.
Ex situ electroplating of the BiFE on the rotating disc electrode (RDE)
Two electroplating solutions were investigated, (A) and (B).
Acetate buffer Bi(III) plating solution. 100 ppm Bi(III) in 1 M acetate buffer (pH 4.5). Plated for 5 mins at −1.0 V in a quiescent solution.LiBr/ HCl Bi(III) plating solution. 0.02 M Bi(III) in 0.5 M LiBr and 1 M HCl. Plated for 60 s at −0.28 V in a quiescent solution.
The RDE was then carefully rinsed before transferring it to the analysis solution.
In-line electroplating of the BiFE on the flow cell working electrode
Two electroplating solutions were investigated, (A) and (B).
Acetate buffer Bi(III) plating solution. 100 ppm Bi in 0.1 M acetate buffer (pH 4.5). Plated for 5 mins at −1.0 V at a flow rate of 37.1 μl/s.LiBr/ HCl Bi(III) plating solution. 0.02 M Bi(III) in 0.5 M LiBr and 1 M HCl. Plated for 120 s at −0.28 V at a flow rate of 37.1 μl/s.
After formation of the BiFE, the plating flow stream was flushed with water and switched to the analysis stream.
Bismuth film cleaning procedure
The Bi(III) film was cleaned of the remaining adsorbed complexes by holding the electrode at a potential of −1.3 V for 30 s in 0.1 M HCl.
Bismuth film removal
Bi(III) films were removed by applying a conditioning potential at +0.3 V, −0.5 V and 0 V for 2 × 50 s in a flowing solution of 0.1 M HNO3.
In-line plating of MFE on the flow cell working electrode
The Hg plating solution was 200 ppm in 0.1 M HNO3. Plated for 120 s at −1 V at a flow rate of 9 μl/s. After film formation the cell was flushed with water and switched to the analysis stream.
Mercury film cleaning procedure
The film was cleaned at a potential of −1.4 V for 10 s in 0.1 M HNO3.
Mercury film removal
Films were removed at +0.8 V in a flowing solution of 0.1 M HNO3 for 60 s.
Results and discussion
The in-line prepared BiFE was assessed for its suitability in performing CAdSV of Co(II) and Ni(II). Various parameters, such as flow rates, solution composition, plating parameters and square-wave conditions, were studied to investigate the electrochemical behaviour of the Bi(III) films prepared on the injection-moulded carbon fibre electrodes.
Investigation of the effect of flow rate during the plating and accumulation stages
Using the acetate buffer-based plating solution, i.e. the 100 ppm Bi(III) in 0.1 M acetate buffer, the effect of varying the flow rate during BiFE plating procedure was examined at 20, 25 and 37 μl/s by varying the height of the reservoir in the gravity-fed flow system (with sample conditions as given in Fig. 2). The maximum response was obtained for the stripping peaks at a flow rate of 37 μl/ s and so this rate was subsequently employed in this study.
Fig. 2Influence of flow rate during the accumulation stage on the Ni(II) and Co(II) CAdSV peaks, where the flow rates are: (a) 5.8, (b) 15.7, (c) 25.6, (d) 37.1, (e) 42.9 and (f) 55.5 μl/s respectively. Carried out with a 10 ppb sample of Co(II) and Ni(II) in 0.1 M ammonium buffer (pH 9.2) with 0.1 mM DMG and 0.5 M NaNO2. SW conditions: Conditioning potential −1.3 V (10 s), deposition potential −0.8 V (120 s), equilibration time 5 s, pulse frequency 15 Hz, pulse amplitude 40 mV and step potential 4 mV. Scan: initial −0.8 V to end potential −1.3 V. Data were smoothed with a level-2 Savitzky–Golay filter
The effect of varying the flow rate during the accumulation stage on the peak heights of Co(II) and Ni(II) was studied over the range of 5.8–55.5 μl/s; the results are depicted in Fig. 2. The response for Co(II) increased linearly with increasing flow rate with a slope of −0.036 μAs/ μl and an intercept of −1.26 μA with a correlation coefficient (R2 = 0.96), whereas that for Ni(II) did not show significant variation. A flow rate of 37 μl/s was selected, as this gave an acceptable response without adverse peak broadening and was compatible with the BiFE plating step flow rate, thereby simplifying the instrumentation.
Effect of composition of the BiFE plating solution
Reported methods for AdSV of Co(II) and Ni(II) have generally employed acetate buffer (pH 4.5) containing 100 mg/l Bi(III) [15, 24, 26, 28]. However, Królicka et al. [29, 30] suggested the use of a bromide-modified plating solution. Hence, both plating solutions were assessed using the RDE and flow cell setups. Figure 3 depicts the differing performances of the two plating solutions. The LiBr-containing plating solution gave the best stripping responses on both the RDE and the flow cell and was therefore used for subsequent studies.
Fig. 3Comparison of different Bi(III) plating solutions. a Effect of (a) 100 ppm Bi(III) in acetate buffer (1 M, pH 4.5) and (b) 0.02 M Bi(III) in 0.5 M LiBr- and 1M HCl-based preplating solutions on RDE at 500 rpm. b Effect of (c) the acetate buffer (0.1 M, pH 4.5) and (d) LiBr/HCl preplating solutions on the flow cell. c Comparision of acetate buffer pH 4.5 Bi(III) preplating solution on (e) flow cell (0.1 M) and (f) RDE (1 M). d Comparison of LiBr preplating solution on (g) flow cell and (h) RDE. Carried out with a 10 ppb sample of Co(II) and Ni(II) in 0.1 M ammonium buffer (pH 9.2) with 0.1 mM DMG and 0.5 M NaNO2. SW conditions: Conditioning potential −1.3 V (10 s), deposition potential −0.8 V (120 s), equilibration time 10 s, pulse frequency 25 Hz, pulse amplitude 50 mV and step potential 5 mV (except trace (e) which was at 15 Hz, 40 and 4 mV respectively). Scan: initial −0.8 V to end potential −1.3 V
Effect of solution parameters on the system behaviour
Buffer concentration The effect of changing the ammonium buffer concentration over the range 0–0.5 M is shown in Fig. 4a. Increasing the buffer concentration over the range 0–0.1 M had a marked effect on the height of the Co(II) peak, but increasing the concentration further did not improve the peak height due to the increased background signal. As Co(II) and Ni(II) were sufficiently resolved at 0.1 M, this concentration was selected for all further experiments.
Fig. 4Effect of solution parameters on the CAdSV behaviour of a 10 ppb Co(II) and Ni(II) sample: (a) ammonium buffer (pH 9.2) concentration over the range 0–0.5 M, (b) DMG concentration over the range 0–0.35 M and (c) nitrite concentration over the range 0–0.8 M. Supporting electrolyte conditions were: (b and c) 0.1 M ammonium buffer (pH 9.2), (a and c) 0.1 mM DMG and (a and b) 0.5M NaNO2. SW conditions were: conditioning potential −1.3 V (10 s), deposition potential −0.8 V (120 s), equilibration time 10 s, pulse frequency 25 Hz, pulse amplitude 40 mV (except c: 50 mV) and step potential 4 mV (except c: 5 mV). Scan: initial −0.8 V to end potential −1.3 V
Effect of DMG concentration Figure 4b shows the effect of increasing DMG concentration, over the range 0.01–0.35 mM, on the peak current. The Co(II) signal increases almost linearly with increasing DMG concentrations, whereas the signal for Ni(II) steadily decreased to 0.2 mM whereupon it levelled off. The signal for the Co(II) peak shifted to a slightly more negative potential (−4 mV) than that for Ni(II) with increasing concentration. A concentration of 0.3 mM was employed in the subsequent studies.
Effect of nitrite concentration Figure 4c shows the influence of nitrite concentration on the sample signal. The Co(II) signal shows a rapid increase in signal to 0.3 M and a slower increase thereafter, whereas the Ni(II) peak height remained level to a concentration of 0.6 M. Concentrations of greater than 0.6 M lead to increased background current and so a nitrite concentration of 0.5 M was subsequently employed. Figure 5 shows the results obtained from the addition of nitrite to the sample system. Addition of nitrite to separate samples of Ni(II) and Co(II) results in increases in response for both metals, Fig. 5a and b, whereas the addition of nitrite to a mixture of 10 ppb Co(II) and Ni(II) results in an almost unchanged Ni(II) peak height whilst that for Co(II) has increased elevenfold, Fig. 5c. Figure 5d shows the reduction (16.6%) of the Co(II) peak height upon the addition of 10 ppb Ni(II), while Fig. 5e displays the reduction (51.9%) of the Ni(II) peak height upon the addition of 10 ppb Co(II).
Fig. 5a Effect of nitrite on stripping peak for 10 ppb Ni(II) when (a) no NaNO2 is present; (b) 0.5 M NaNO2 is present. b Effect of nitrite on stripping peak for 10 ppb Co(II) when (c) no NaNO2 is present; (d) 0.5 M NaNO2 is present. c Effect of nitrite on the stripping peaks for 10 ppb Co(II) and Ni(II) when (e) no NaNO2 is present; (f) 0.5 M NaNO2 is present. d Effect of addition of 10 ppb Ni(II) on the peak height for 10 ppb Co(II) in the presence of 0.5 M NaNO2 when (h) Co(II) only is present and (g) both Co(II) and Ni(II) are present. e Effect of addition of 10 ppb Co(II) on the peak height for 10 ppb Ni(II) in the presence of 0.5 M NaNO2 when (j) Ni(II) only is present and (k) Co(II) and Ni(II) are present. Carried out in 0.1 M ammonium buffer (pH 9.2) with 0.1 mM DMG. SW conditions: conditioning potential −1.3 V (10 s), deposition potential −0.8 V (120 s), equilibration time 5 s, pulse frequency 25 Hz, pulse amplitude 50 mV and step potential 5 mV. Scan: initial −0.8 V to end potential −1.3 V
Effect of varying the square-wave (SW) stripping parameters
Frequency The effect of frequency was studied in the range 12.5–100 Hz, Fig. 6a depicts the results obtained. The peak heights for both metals increased with increasing SW frequency (as expected from increasing the effective scan rate) and shifted towards more negative values. However, increasing the frequency leads to insufficient time for the background current to decay and leads to an increase in the contribution to the signal and hence a distortion in peak shape. A frequency of 25 Hz was generally the maximum employed.
Fig. 6Effect of varying the SW stripping parameters on the CAdSV peaks of a 10 ppb Co(II) and Ni(II) sample. The variables are (a) frequency (12.5–100 Hz), (b) step potential (0.15–7 mV), (c) amplitude (5–70 mV), (d) deposition potential (−0.3 to −0.9 V), (e) deposition time (30–210 s). Supporting electrolyte: 0.1 M ammonium buffer (pH 9.2) with 0.1 mM DMG and 0.5 M NaNO2. SW conditions were (unless being investigated as above): conditioning potential −1.3 V (10 s), deposition potential −0.8 V (120 s), equilibration time 5 s, frequency 25 Hz, pulse amplitude 50 mV (except b and e: 40 mV) and step potential 5 mV (except e: 4 mV). Scan: initial −0.8 V to end potential −1.3 V
Step potential Figure 6b shows the effect of the step potential on the peak heights. The response for Co(II) rose rapidly with rising step potentials (i.e. with an increase in effective scan rate) up to 3 mV and thereafter levelled off, whereas the response for Ni(II) increased slowly to 7 mV. Peak resolution remained the same across this step potential range, with both peaks shifting towards more negative potentials. Values of 4 or 5 mV were generally found to be ideal for adequate peak definition.
Amplitude The effect of the SW pulse amplitude was studied over the range 5–80 mV. Both peaks shifted to more positive values, see Fig. 6c and they both initially increased rapidly in height (5–20 mV) whereupon the responses for Co(II) and Ni(II) levelled off as a result of the increase in contribution from the background current. A SW pulse amplitude of either 40 or 50 mV was employed.
Deposition potential Figure 6d shows the influence of the deposition potential on the peak currents. At potentials less than −0.4 V the Ni(II) signal is completely suppressed and the Co(II) signal was markedly reduced due to the oxidation of the bismuth film (at around −0.46 V in alkaline solutions). Both peaks gradually increase with potential in the range of −0.4 V to a maximum at −0.8 V and then exhibit a sharp reduction in adsorption at a more negative potential of −0.9 V resulting from further reduction of the analyte complex. A deposition potential of −0.8 V was selected, as this gave the maximum response for both metals.
Accumulation time The influence of the accumulation time on the stripping signal was examined. Figure 6e depicts the response in peak currents for 10 ppb each of Ni(II) and Co(II). For both Co(II) and Ni(II) the response increases with accumulation time up to 150 s for Co(II) and 180 s for Ni (II), after which the response levels off, indicating saturation of the surface. An accumulation time of 120 s was used for subsequent measurements as this gave the best compromise between sensitivity and analysis time.
Analytical characterisation
The simultaneous determination of Co(II) and Ni(II) over the concentration range of 1–20 ppb was investigated. The determination was carried out in 0.1 M ammonium buffer (pH 9.2) with 0.1 mM DMG and 0.5 M NaNO2. SW conditions: conditioning potential −1.3 V (10 s), deposition potential −0.8 V (120 s), equilibration time 5 s, pulse frequency 25 Hz, pulse amplitude 50 mV and step potential 5 mV. Scan: initial −0.8 V to end potential −1.3 V. The BiFE exhibited a well-defined response to changing concentration and the peak current response (y, μA) was linear over the concentration range (x, ppb) of 1–20 ppb, as expressed in the calibration plots for Co(II),y = −11.4x − 85.5 (R2 = 0.96), and Ni(II),y = −1.64x – 7.20 (R2 = 0.92), respectively.
The stability of the BiFE formed on the injection-moulded substrates was investigated by performing replicate runs on the same film. Ten replicate runs gave an average peak height for Ni(II) of −1.05 ± 0.10 × 10−5 A (rsd: 9.6%) and for Co(II) −8.19 ± 0.82 × 10−5 A (rsd: 10.0%).
To compare the working electrode behaviour, AdSV and CAdSV of Co(II) and Ni(II) were also performed at a MFE in the flow cell. The solutions needed to be deaerated before analysis on the MFE. Neither AdSV or CAdSV of Co(II) on a MFE was possible in these particular flow cells due to a significant shift in the stripping peak to an extreme cathodic potential (<−1.4 V vs Ag/AgCl), which led to analytical uncertainty at the edge of the supporting electrolyte potential window. However, AdSV of Ni(II) was possible, as illustrated in Fig. 7. It has been noted previously by this group [57, 58] that the main drawback with MFEs is the difficulty involved in regenerating the active mercury surface after use, necessitating the inclusion of cleaning and mercury film renewal procedures. In contrast to the MFEs, the BiFEs formed in these flow cells enabled the generation of reproducible, quantitative analytical data, due to the enhanced stability of the bismuth films during the cleaning cycles, and the reduced cathodic potential for the cobalt stripping peak.
Fig. 7AdSV of Ni(II) on a mercury film electrode. Here decimal numerals indicate the film number and roman numerals indicate replicate analyses carried out on the same film. Carried out with a 200 ppb sample in 0.1 M ammonium buffer (pH 9.3) with 0.1 mM DMG (deaerated) at a flow rate of 16 μl/s. SW conditions: conditioning potential −1.4 V (10 s), deposition potential −0.5 V (200 s), equilibration time 5 s, pulse frequency 25 Hz, pulse amplitude 10 mV and step potential 4 mV. Scan: −0.5 V to end potential −1.4 V. Data were smoothed with a level-2 Savitzky–Golay filter
Conclusions
The application of a BiFE for the determination of Co(II) and Ni(II) by square-wave CAdSV under flow conditions in a non-deaerated solution has been demonstrated. The injection-moulded carbon-fibre-filled polystyrene electrodes have proven to be suitable supports for the formation of bismuth films. The LiBr-containing plating solution was found to offer superior performance for both the RDE and the flow cell under the alkaline conditions required for CAdSV. The application of a flow stable BiFE in an inexpensive sensor system permits the possibility of autonomous on-site industrial and environmental monitoring. The inherent disposability of the devices coupled with their mercury-free status should advance the scope of the sensor to point-of-care clinical applications.

square-wave adsorptive stripping voltammetry


High-throughput screening of monoclonal antibodies against plant cell wall glycans by hierarchical clustering of their carbohydrate microarray binding profiles

Antibody-producing hybridoma cell lines were created following immunisation with a crude extract of cell wall polymers from the plant Arabidopsis thaliana. In order to rapidly screen the specificities of individual monoclonal antibodies (mAbs), their binding to microarrays containing 50 cell wall glycans immobilized on nitrocellulose was assessed. Hierarchical clustering of microarray binding profiles from newly produced mAbs, together with the profiles for mAbs with previously defined specificities allowed the rapid assignments of mAb binding to antigen classes. mAb specificities were further investigated using subsequent immunochemical and biochemical analyses and two novel mAbs are described in detail. mAb LM13 binds to an arabinanase-sensitive pectic epitope and mAb LM14, binds to an epitope occurring on arabinogalactan-proteins. Both mAbs display novel patterns of recognition of cell walls in plant materials.
Introduction
Carbohydrate microarrays provide a means of rapidly screening the interactions between glycans and other molecules [1–7]. Applications for this technology include the screening of protein–glycan interactions, characterization of carbohydrate-active enzymes and the analysis of the specificities of monoclonal antibodies (mAbs) and carbohydrate-binding modules [8, 9]. mAbs are powerful tools for investigating the biological roles of glycans but producing them is time consuming, labour-intensive and costly. Hybridoma-based mAb production involves the fusion of spleen cells from an immunized animal with myeloma cells. The resulting hybridoma cell lines are both immortal and secrete antibody into the cell supernatant [10]. This procedure is conventionally performed in a one-by-one fashion, such that each animal is immunized with a single antigen and the resulting antibodies are screened for desired specificities using enzyme-linked immunosorbent assays (ELISAs). However ELISA-based screening is low throughput because only a limited amount (∼100 μl) of hybridoma supernatant is available for screening during the initial stages of mAb production and this is typically only sufficient to test each mAb against just one or two antigens. An alternative approach involves ‘shotgun’ immunisation with a mixture of antigens [11]. This potentially results in the simultaneous generation of antibodies with a range of specificities, but the time limiting step then becomes the detailed retrospective screening of their specificities. However, microarrays offer a means of rapidly screening limited amounts of hybridoma supernatant against multiple antigens and therefore greatly increasing throughput in the identification of valuable cell lines. Here we report the use of shotgun immunisation followed by microarray-based screening of hybridoma supernatants in order to produce mAbs against plant cell wall glycans.
Plant cell walls are fibre composites that contain some of the most complex glycans known [12, 13]. In addition to their biological roles, many cell wall components have important industrial applications including as functional food ingredients, pharmaceuticals, nutriceuticals, fibres and increasingly, bio-fuels [14–16]. Cell wall glycans can be broadly grouped into cellulose, hemicelluloses, pectins and glycoproteins [17]. Cellulose microfibrils are cross-linked by hemicelluloses such as xyloglucans, xylans and mixed linkage glucans forming a tough load-bearing matrix which is embedded in pectic polysaccharides [18]. Pectins are the most complex and heterogeneous family of cell wall glycans and are comprised of a series of galacturonic acid-rich polymers including homogalacturonan (HG), rhamnogalacturonan I (RGI), rhamnogalacturonan-II (RG-II) and xylogalacturonans (XG). In addition, structurally complex arabinan, galactan and arabinogalactan polymers may be present as side chains to the galacturonan-rich backbone domains [18]. The protein moieties of cell wall glycoproteins are often rich in hydroxyproline (Hyp) and these polymers, which are collectively referred to as Hyp-rich glycoproteins (HRGPs), include the extensins and arabinogalactan-proteins (AGPs) [19–21].
The fine structures and relative amounts of cell wall components vary greatly not only among plants, but also between organs, tissues, cells, and even between different micro-domains within a single cell wall. This complexity and heterogeneity presents a major barrier to detailed analysis and our understanding of many aspects of plant cell wall structure and function is far from complete. Several mAbs with specificities for diverse plant cell wall components have been developed and these are powerful tools for the analysis of cell walls [22]. However, the repertoire of mAbs currently available covers only a small proportion of the glycan structures that have been identified and there is a pressing need for a wider range of mAbs to facilitate the further characterization of cell walls. We have developed a method of rapidly screening mAb specificities using microarrays of cell wall glycans including pectins with different degrees and patterns of methyl-esterification, pectic side chains (e.g. arabinan and galactan), hemicelluloses (e.g. xylans, mannans, and xyloglucans), HRGPs (e.g. AGP-rich gums), β-linked glucans (e.g. (1→3)(1→4)-β-glucan and (1→3)-β-glucan) and celluloses (e.g. hydroxylethyl cellulose and carboxylmethyl cellulose). A further 14 samples were sequentially extracted from A. thaliana using CDTA and NaOH which are known to predominately solublize pectic polymers and hemicelluloses respectively [23].
Experimental procedures
Immunisation and generation of monoclonal hybridomas
Cell wall polymers were isolated from 6 week old wild type A. thaliana plants, ecotype Col-0, grown in soil at 22°C with cycles of 10 h of light/14 h of darkness. One g dry weight of a mixture of leaves, stems and roots was homogenized to a fine powder in liquid nitrogen. The homogenate was incubated with 20 ml 50 mM 1,2-Diaminocyclohexanetetraacetic acid (CDTA; pH 7.5) for 3 h at 18°C and centrifuged for 20 min at 4,400 rpm. The supernatant was collected and dialyzed extensively against deionized water (dH2O) in dialysis tubing (6–8,000 kDa molecular weight cut off) to remove low molecular weight molecules and freeze dried. The material was dissolved in phosphate-buffered saline (PBS) to generate the immunogen. Rats were used for antibody production so as to make subsequent comparisons with existing mAbs, most of which were produced in rats, as valid as possible. The immunization of rats, hybridoma preparation and cloning procedures were as described previously [24]. Briefly, two male Wistar rats were each injected subcutaneously with 250 μl of an emulsion of the isolated cell wall material at 1 mg/ml in PBS with an equal volume Freund’s complete adjuvant on day 0. On days 40 and 79 the injections were repeated using incomplete adjuvant. Tail bleeds were taken 10 days after injections to assess the immune response. On day 198 a pre-fusion boost was given to the selected rat and 3 days later, the spleen was removed and lymphocytes were isolated and fused with rat myeloma cell line IR983F [25] using standard polyethylene glycol fusion of lymphocytes and myeloma cells. Hybridoma lines were initially screened by ELISA with the immunogen coated onto microtitre plates (MaxiSorp, Nunc, Roskilde, Denmark) at 50 μg/ml.
Previously described monoclonal antibodies
Arrays were probed with 23 mAbs with previously described specificities and details of these are provided in Table 1. All were rat antibodies except mAbs CCRC-M1, BS-400-02, BS-400-03, and BS-400-04 which were produced in mice and PAM1 which was produced by phage display.
Table 1Previously characterized monoclonal antibodies used to probe glycan arrays. HG, homogalacturonanSpecificityNameRef.Un-esterified HGPAM1[31]Un-esterified/Calcium ion cross-linked HG2F4[34]Partially methyl-esterified HGJIM5[33]Partially methyl-esterified HGJIM7[33](1→4)-β-galactanLM5[37](1→5)-α-arabinanLM6[24]Fucosylated xyloglucanCCRC-M1[38]Non-fucosylated xyloglucanLM15(1→4)-β-mannan/galacto-(1→4)-β-mannanBS-400-4[39](1→4)-β-xylanLM10[40](1→3)(1→4)-β-glucanBS 400-3[41](1→3)-β-glucanBS 400-2[32](1→4)-β-xylan/arabinoxylanLM11[40]Arabinogalactan-proteinJIM4[43]Arabinogalactan-proteinLM2[42]Arabinogalactan-proteinMAC207[43]Arabinogalactan-proteinJIM8[44]Arabinogalactan-proteinJIM13[43]Arabinogalactan-proteinJIM16[43]Arabinogalactan-proteinJIM14[43]ExtensinLM1[45]ExtensinJIM19[46]ExtensinJIM20[46]
Glycan samples used on the array
Details of the 50 defined glycans used are provided in Table 2. Most glycans were dissolved in dH2O. Arabinoxylan and glucuronoxylan were prepared by boiling in dH2O for 10 min. and then standing for 3 h at 18°C before use. Glucomannan was prepared by wetting with 95% ethanol followed by addition of dH2O. The mixture was heated to boiling point and stirred for 20 min until dissolved. Pachyman was prepared by dissolution in a minimal volume of 10% (w/v) sodium hydroxide followed by neutralization with acetic acid. 14 samples on the arrays were cell wall polymers extracted from A. thaliana organs listed in Table 2 using CDTA and 4 M NaOH. Fifty milligrams (fresh weight) of each organ collected from at least four separate plants were homogenized to a fine powder prior to adding 300 μl of 50 mM CDTA (pH 7.5). After incubating with rotation for 4 h at 20°C, the extracts were centrifuged at 4,400 rpm for 10 min and the supernatants (‘CDTA extracts’) removed. Pellets were resuspended in 300 μl of 4 M NaOH and samples were incubated with rotation for 4 h at 20°C prior to centrifugation at 4,400 rpm for 10 min. Supernatants were ‘NaOH extracts’.
Table 2Samples included on the glycan arraysAlphanumerical codesSamplesA1Arabinan (sugar beet)B1Pectin (apple)C1Galactan (lupin)D1Homogalacturonan (sugar beet)E1Pectin (lime) B15F1Pectin (lime) B43G1Pectin (lime) B71H1Pectin (lime) 96A2Pectin (lime) F11B2Pectin (lime) F19C2Pectin (lime) F43D2Pectin (lime) F76E2Pectin (lime) P16F2Pectin (lime) P24G2Pectin (lime) P32H2Pectin (lime) P41A3Pectin (lime) P46B3Pectin (lime) P60C3Pectin (lime) P76D3RGI (soybean)E3RGII (A. thaliana)F3Xylogalacturonan (pea)G3MHR I (apple)H3MHR II (carrot)A4MHR III (potato)B4MHR HS1 (apple)C4MHR HS2 (apple)D4Xylogalacturonan (apple)E4AGP (P. patens)F4Seed mucilage (A. thaliana)G4Xyloglucan/mannan (tomato)H4Glucomannan (konjac)A5Gum (guar)B5Gum (locust bean)C5Gum arabic (acacia)D5Gum (karaya)E5Gum (tragacanth)F5AGP (larch)G5Arabinoxylan (wheat)H5β(1-3),(1-4)-glucan (lichenan)A6Mannan (ivory nut)B6Xyloglucan (tamarind)C6Glucuronoarabinoxylan (maize)D6Hydroxyethyl celluloseE6β(1-4)-glucan (avicel)F6Carboxymethyl celluloseG6Alginic acidH6β(1-3),(1-6)-glucan (laminarin)A7β(1-3)-glucan (pachyman)B7β(1-4),(1-6)-glucan (pullulan)C7CDTA extract (A. thaliana flowers)D7CDTA extract (A. thaliana siliques)E7CDTA extract (A. thaliana stem top)F7CDTA extract (A. thaliana stem middle)G7CDTA extract (A. thaliana stem base)H7CDTA extract (A. thaliana leaves)A8CDTA extract (A. thaliana roots)B8NaOH extract (A. thaliana flowers)C8NaOH extract (A. thaliana siliques)D8NaOH extract (A. thaliana stem top)E8NaOH extract (A. thaliana stem middle)F8NaOH extract (A. thaliana stem base)G8NaOH extract (A. thaliana leaves)H8NaOH extract (A. thaliana roots)Alphanumerical codes refer to the position of samples on arrays. Source organisms are in parenthesesRGI Rhamnogalcturonan I; RGII rhamnogalacturonan II; MHR modified hairy region; AGP arabinogalactan-protein
Post-printing modification of glycans
Glycan samples on selected arrays were modified in situ after printing by enzymatic digestion. For the data in Fig. 6 selected arrays were digested with endo-α(1-5)-L-arabinanase or endo-β(1-4-)-galactanase (both from Aspergillus niger, Megazyme (Bray, Ireland) used at 1 U/ml in 200 mM sodium acetate pH 4.0.
Printing of arrays
Glycans were applied to nitrocellulose membrane (0.45 μm pore size, Schleicher and Schuell, Dassel, Germany) at two concentrations (0.2 and 0.04 mg/ml) and in duplicate such that each sample was represented by four spots. CDTA and NaOH extracted A. thaliana material was printed as extracted and as a five fold dilution, also in duplicate. Printing was performed using a microarray robot (Microgrid II, Genomic Solutions, Ann Arbor, MI, USA) equipped with split pins (MicroSpot 2500, Genomic Solutions). Pins were washed twice in dH2O after deposition of each sample.
Probing of arrays
Arrays were blocked by incubation for 1 h in PBS (140 mM NaCl, 2.7 mM KCl,10 mM Na2HPO4, 1.7 mM KH2PO4, pH 7.5) containing 5% w/v low fat milk powder (5%MPBS). Arrays were then probed for 2 h with antibodies diluted in 5%MPBS. All antibodies were used as 1/10 dilutions except CCRC-M1 which was used at 1/50, and BS-400-2, BS-400-3 and BS-400-4 which were used at 1/200. After washing with PBS, arrays were incubated for 2 h in either anti-rat or anti-mouse secondary antibody conjugated to alkaline phosphatase (Sigma, Poole, UK) diluted 1/5000 in 5%MPBS. After washing in PBS, arrays were developed using a substrate containing 5-bromo,4-chloro,3-indolylphosphate (BCIP) and nitroblue tetrazolium (NBT) in BCIP/NBT buffer (100 mM NaCl, 5 mM MgCl2, 100 mM diethanolamine, pH 9.5).
Scanning and analysis
Arrays were scanned, converted to 16 bit grey-scale TIFFs, transformed to negative images and uploaded into ImaGene 6.0 microarray analysis software (BioDiscovery, El Segundo, CA, USA). Semi-automatic gridding was used to create an analysis area for each spot, and a 5-pixel zone around each spot was used for calculation of local background signals. Individual spot signals were defined as the mean pixel value within each spot area (red zone in Fig. 1d) minus the median pixel value in the surrounding local background area (green zone in Fig. 1d). The mean of the four individual spot signals for each sample was defined as the ‘mean sample value’ (MSV) and ‘total mean sample values’ (TMSVs) were the means of MSVs from three separate experiments. Selected data sets (TMSVs) were used to generate heatmaps (Figs. 2b and 5b) using online heatmapper software (http://bbc.botany.utoronto.ca/ntools/cgi-bin/ntools_heatmapper.cgi). For each antibody in the heatmap, the maximal TMSV was set to 100% and all other values within that data set adjusted accordingly. A cut off of 5% of the maximal TMSV was imposed. Hierarchical cluster analysis was performed on TMSVs obtained for 23 previously characterized antibodies and 7 newly produced ones using Epclust software (http://ep.ebi.ac.uk/EP/EPCLUST/). Clustering was based on correlation measure based distance and average linkage.
Fig. 1Microarrays of plant cell wall glycans. a, b A library of 50 glycan polymers were spotted onto nitrocellulose membranes in an area 22 mm × 22 mm. Arrays included pectins, HRGPs, hemicelluloses and glucans (samples, A1–B7) and the position of these classes of polymer on the arrays are indicated by colour coding in (b). Fourteen samples extracted from Arabidopsis thaliana using either CDTA (samples C7–A8) or NaOH (samples B8–H8), were also included. Details of the samples are provided in Table 2. c Each sample was printed at the two concentrations indicated and in duplicate. d Microarray analysis software (ImaGene 6.0) was used to quantify spot signals and individual spot signals were defined as the mean pixel value within each spot area (red zone) minus the median pixel value in the surrounding local background area (green zone). e–g Two pairs of three arrays were printed and probed with the anti-homogalacturonan mAb JIM5 on three separate occasions. The mean sample values (MSVs) for each pair were quantified and plotted against MSVs of an array produced on a different occasion. These are presented as scatter plots, with the R2-values for the plots indicatedFig. 2Probing of cell wall glycan arrays with mAbs with known specificities. a Glycan arrays were probed with 23 mAbs with previously defined specificities and 5 representative examples are shown: PAM1 (anti-homogalacturonan); BS-400-2 (anti-(1→3)-β-glucan; LM5 (anti-(1→4)-β-galactan); JIM13 (anti-arabinogalactan-protein); JIM5 (anti-homogalacturonan). A control array was probed with secondary antibody, but no primary mAb. Arrays were scanned and converted into 16 bit greyscale TIFFs. b Heatmap showing the total mean sample values (TMSVs) for the binding of each mAb to each sample. The maximal TMSV for each mAb was set to 100 and all other values were adjusted accordingly. A cut off of 5% of the maximal TMSV was imposed and all values at, or below this are represented as black boxes
Indirect immunofluorescence labeling of plant materials
Resin embedded and fresh plant material was sectioned and labelled with antibodies as described previously [26, 27]. LM14 and LM13 hybridoma supernatants were used as 1/10 dilutions in 5%MPBS. Sections were counterstained for cellulose with Calcofluor white (fluorescent brightener 28, Sigma, Poole, UK) used as a 0.005% aqueous solution.
Competitive-inhibition ELISA assays
The ability of arabinose, galactose, rhamnose and oligoarabinosides to inhibit the binding of LM14 and LM13 to the immuogen in ELISAs was assessed as described previously [28].
Sugar composition analysis of modified pectic hairy regions (MHRs)
Sugar composition was determined using methanolysis as described previously [29]. MHRs were treated with 2 N HCl in dry methanol for 16 h at 80°C, followed by 1 h of 2 M CF3CO2H (TFA) at 121°C. The released sugars were analysed using high-performance anion exchange chromatography (HPAEC) with pulsed amperometric detection (PAD) as described previously [30].
Results
Production of microarrays of plant cell wall polymers
The layout of the arrays is shown in Fig. 1 and colour coding indicates the distribution on the arrays of different classes of cell wall polymer (Fig. 1a,b). A detailed list of the samples used is provided in Table 2. Each sample was represented on the arrays by four spots (two concentrations, printed in duplicate) (Fig. 1c) and microarray analysis software was used to quantify background-corrected spot signals (Fig. 1d). As shown by the arrays in Figs. 2a and 3, the consistency of the printing and probing replicates within arrays was high. However, reproducibility between arrays was also tested. A total of six arrays (3 × 2 pairs) were printed and probed with the anti-HG mAb JIM5 on three separate occasions. The MSVs were quantified and the values for each array were plotted against those from an array produced on a different occasion. The R2-values obtained for these plots were all >0.98, indicating a high degree of reproducibility between arrays (Fig. 1e–g).
Fig. 3Probing of cell wall glycan arrays with newly produced mAbs. Glycan arrays were probed with mono- or multiclonal antibodies generated following shotgun immunisation with plant cell wall polymers. Four representative examples are shown. Arrays were scanned and converted into 16 bit greyscale TIFFs
Probing of plant cell wall glycan arrays
Arrays were probed with a range of mAbs with previously defined specificities for epitopes occurring on the major classes of cell wall polymers. Details of the mAbs used are provided in Table 1. A total of 23 mAbs were tested and 5 representative examples are shown in Fig. 2a. The TMSVs for the binding of these mAbs to each sample (derived from three independent experiments) are shown as a heatmap in Fig. 2b, where spot signals are correlated to colour intensity. mAb binding profiles were in good agreement with the published results for the antibodies tested. For example, mAb PAM1 has specificity for HG with a low degree of methyl-esterification (DE) [31] and consistent with this, PAM1 bound to lime pectin samples with DEs of 11–19% (samples E1, A2 and B2). PAM1 did not bind above background to other pectin samples with higher DEs, or to any other cell wall glycans. mAb BS-400-2 is specific for (1→3)-β-glucan [32] and, of the defined samples, only bound to this polymer (sample A7) on the arrays. (1→3)-β-glucan is known to be extractable from plant cell walls using NaOH and consistent with this, BS-400-2 also bound to weakly to NaOH-solubilized extracts from A. thaliana (samples B8-H8). As expected, the epitopes recognized by some mAbs were detected on multiple samples and this was the case for mAbs LM5 (anti-(1→4)-β-galactan), JIM13 (anti-AGP) and JIM5. However, the binding profiles of these mAbs were also consistent with their known specificities. For example, JIM5 bound with greatest avidity to pectin samples that contained abundant HG domains (such as the lime pectins E1-C3), but not to pectic fragments lacking HG (such as the arabinan or galactan samples A1 and C1).
To generate a new series of cell wall-directed mAbs, rats were immunized with an immunogen consisting of a crude extract of cell wall material solubilized from A. thaliana using CDTA. 13 cell lines were selected by an initial ELISA analysis against the immunogen and these were subsequently probed against the cell wall polymer arrays and four representative examples are shown in Fig. 3. Antibodies that bound with very low avidity, or appeared to have identical binding profiles to previously generated antibodies were not selected for further analysis whilst the data for the remaining seven were subjected to hierarchical cluster analysis.
Hierarchical clustering of antibody binding profiles
Cluster analysis was used to rapidly obtain information about the specificities of 7 new antibodies based on the similarly of their binding profiles on the arrays to 23 previously characterized mAbs (Fig. 4). Clustering indicated that most of the antibodies tested separated into 4 broad clusters: HG-binders; pectic side chain-binders; HRGP-binders and hemicellulose-binders. Three antibodies with specificity for (1→3)(1→4)-β-glucan, (1→3)-β-glucan and (1→4)-β-mannan did not cluster with any other mAbs within the 0.55 (distance measurement units) cut off imposed. These broad clusters were in agreement with the previously defined specificities of the mAbs tested, and this was also the case within sub-clusters. For example, within the HG-binders cluster, mAbs JIM5 and JIM7, which both bind to partially methyl-esterified HG [33], clustered closely, whilst mAbs 2F4 and PAM1, which both bind to non-methyl-esterified HG also clustered closely [31, 34]. Similarly, within the hemicellulose-binders, the anti-xylan mAbs LM10 and LM11 formed a sub-cluster, as did the two anti-xyloglucan mAbs CCRC-M1 and LM15. One of the new mAbs (1H12, designated LM13 when cloned) was grouped with the pectic side chain-binders LM5 (anti-(1→4)-β-galactan) and LM6 (anti-(1→5)-α-arabinan). However, most of the new antibodies tested were grouped in the HRGP-binders cluster and within this, clustered most closely with the anti-AGP mAbs MAC207, JIM4 and JIM16. One of the new mAbs within this group (1B3, designated LM14 when cloned) was, together with LM13 subject to more detailed epitope characterization in order to further explore the evidence gained from the clustering analysis.
Fig. 4Cluster analysis of antibody binding profiles to arrays. 23 mAbs with previously defined specificities, and 7 newly produced antibodies (red diamonds in black boxes) were analyzed using Epclust software. Hierarchical clustering was performed using clustered correlation measure based distance and average linkage. The binding profiles of all the mAbs analyzed clustered into four broad groups: 1 homogalacturonan-binders; 2 pectic side chain-binders; 3 HRGP-binders; and 4 hemicellulose-binders. Three mAbs were non-clustering outliers (NCO). The specificities of the 23 previously defined mAbs are listed in Table 1
Detailed characterization of the epitopes recognized by LM14 and LM13
Cluster analysis of LM13 and LM14 suggested that the epitopes recognized by these mAbs may contain neutral sugars such as arabinose and galactose, that are abundant in the side chains of both pectins and HRGPs. The glycan arrays contained five 5 samples of branched (or ‘hairy’) regions of pectic polymers modified by enzymatic digestion (‘modified hairy regions’ or MHRs) that were rich in neutral sugars and for which monosaccharide composition was determined (Fig. 5). The binding of LM14 and LM13 to these samples was therefore of particular interest. LM14 bound only to an MHR from carrot (sample H3), whilst LM13 bound to MHRs from potato and apple (samples G3, A4 and B4). It was noteworthy that the two MHR samples to which LM13 did not bind (samples H3 and C4) had the lowest arabinose content, suggesting that the epitope recognized by this mAb may contain arabinose. In order to explore this possibility further, the sensitivity of the LM13 epitope to arabinanase digestion was investigated. To do this, cell wall glycan microarrays were digested with arabinanase prior to probing with mAbs, and the TMSVs obtained with and without digestion were compared. These data are presented as scatter plots (Fig. 6a–d) in which signals from enzyme treated arrays were plotted against signals from untreated arrays. As a positive control, arabinanase digested and untreated arrays were probed with the anti-(1→5)-α-arabinan mAb LM6 [24]. As expected, the binding of LM6 to all samples was reduced to some extent by arabinanase digestion (Fig. 6a). However, the effect of arabinanase digestion on LM13 binding was even more pronounced than for LM6 and LM13 binding to all samples on the arrays was essentially abolished (Fig. 6b). This suggested that the epitope recognized by LM13 either contained arabinan and was degraded by arabinanase, or that the epitope was attached to arabinan and was released from arrays by arabinanase. In contrast, LM14 binding to all samples was essentially unaffected by arabinanase digestion (Fig. 6c). The effect of galactanase digestion on mAb binding to arrayed samples was also assessed. LM13 binding was reduced to some extent by galactanase digestion, but the reduction was far less for than arabinanase digestion (Fig. 6d). LM14 binding to all samples was unaffected by galactanase digestion, and the ability of this enzyme to degrade galactan was indicated by the fact that the binding of the anti-(1→4)-β-galactan mAb LM5 was essentially abolished by galactanase digestion (data not shown). The epitope recognized by LM13 was further characterized using competitive inhibition ELISA (ciELISA) assays (Fig. 6e). The binding of LM13 to immobilized arabinan was inhibited to 50% of maximal binding by approximately 450 μg/ml of both arabino-α1,5-octaose and arabino-α1,5-pentaose, but oligoarabinosides of lower degrees of polymerisation and rhamnose, galactose and glucuronic acid failed to inhibit binding at the highest concentration tested of 1 mg/ml;. The binding of LM14 to immobilized antigen was not inhibited by any of the haptens tested at concentrations up to 1 mg/ml (data not shown).
Fig. 5Binding of LM13 and LM14 to samples to pectic modified hairy regions (MHRs). a Monosaccharide composition of 5 pectin modified hairy regions (MHRs) from apple (samples G3, B4 and C4), carrot (sample H3), potato (sample A4). b TMSVs for the binding of mAbs LM14 and LM13 to these samples. The maximal TMSV for LM13 and LM14 (to all samples on the array) was set to 100 and all other values adjusted accordingly. A cut off of 5% of the maximal TMSV was imposed and all values at, or below this are represented as black boxesFig. 6The epitope recognized by LM13 is arabinanase sensitive. a–d Scatter plots showing the effect of galactanse and or arabinanase on the binding of LM13, LM14 and LM6 (anti-(1→5)-α-arabinan) to glycan arrays. Arrays were incubated in enzymes after blocking but before probing. TMSVs were determined and plotted against each other. Spots that lie on the dotted line indicate that enzyme digestion had no effect on mAb binding, whereas spots that lie below the dotted line indicate where enzyme digestion reduced mAb binding. e The binding of LM13 to oligoarabinosides was tested by competitive inhibition ELISA, in which arabinan was the immobilized polymer. Oligoarabinosides with degrees of polymerization less than five failed to inhibit LM13 binding, even when used at 1 mg/ml. Both arabino-α1,5-octaose and arabino-α1,5-pentaose inhibited the binding of LM13 by 50% of maximal binding when used at approximately 450 μg/ml
Cluster analysis suggested that LM13 was most similar to that of mAbs with specificity for pectic side chains, whilst the binding of LM14 was most similar to that of anti-AGP mAbs. AGPs can be distinguished from pectic polymers using SDS-PAGE because whereas AGPs resolve as characteristic smears, pectic polymers typically do not enter gels [35]. Western blots were prepared using the same A. thaliana CDTA-extracted material that was used for immunisation and probed with LM13 or LM14 (Fig. 7). LM13 did not bind to material on these blots whereas LM14 bound to a high molecular weight smear (∼70–200 kDa) that is characteristic of AGPs.
Fig. 7Immunoblot of monoclonal antibodies LM13 and LM14. The CDTA-soluble Arabidopsis thaliana fraction that was used as the immunogen was separated by SDS-PAGE with 30 μg protein loading per lane. A Western blot of this material was probed with LM13 and LM14. LM13 did not bind to the blot but LM14-probing produced a smear in the region of 70–200 kDa, which is characteristic of AGPs
Immunolocalization of the LM14 and LM13 epitopes
In order to further investigate the epitopes recognized by LM13 and LM14 in planta a range of plant materials were probed with these mAbs using indirect immunofluorescence labelling. In transverse resin-embedded sections of A. thaliana inflorescence stems, LM13 bound specifically to epidermal cells and most strongly to the tangential cell walls (Fig. 8a). Plant cell wall glycans may be altered during extraction and/or immobilization such that epitopes are changed from their native state in planta. In order to test if this was a factor in determining the arabinase sensitivity of the LM13 epitope, sections of A. thaliana stems were also digested with arabinanase prior to probing with LM13. As shown by comparison of Fig. 8a,b, the binding of LM13 to A. thaliana stem epidermal cell walls was abolished following arabinanase digestion indicating that the epitope-structure immobilised on arrays recognized by LM13 reflected that observed in planta. In contrast to LM13, LM14 bound to some extent to all cell walls in an equivalent section through an A. thaliana stem (Fig. 8d). Labelling of the surface of intact roots of A. thaliana seedlings indicated that LM13 bound very weakly whereas LM14 bound strongly to the surface of root hairs (Fig. 8e and f). In transverse sections through tobacco stems, the LM13 epitope was associated with cell walls adjacent to metaxylem cells of the vascular tissue—a distinctly different labelling pattern to that observed in A. thaliana stems (Fig. 8g). In contrast, LM14 did not label any cell walls in tobacco stems (Fig. 8h).
Fig. 8Indirect immunofluorescence labelling of plant material with LM14 and LM13. LM13 bound strongly to inner and outer radial walls of epidermal cells in transverse sections of A. thaliana inflorescence stem sections. The section is counterstained with calcofluor (blue) that labels β-linked glucans. b LM13 binding to A. thaliana inflorescence stem sections when sections were treated with arabinanase. Arrowheads in a and b indicate the epidermal surface. c Control in which an equivalent stem to that in a was labelled with the absence of primary antibody. d, An equivalent section to that shown in a labelled with LM14 which bound to all cell walls. e LM13 bound very weakly to the intact surface of the roots of A. thaliana seedlings. f LM14 bound strongly to the surface of root hairs (arrowheads) of Arabidopsis seedling roots. g In transverse sections through tobacco stem LM13 bound strongly to cell walls adjacent to files of metaxylem cells (mx). h LM14 did not bind to equivalent sections through tobacco stems. Scale bars = 50 μm for a–d, g and h; 20 μm for g inset, 50 μm for e and f
Discussion
The work presented here demonstrates the potential of glycan microarrays for overcoming a major bottleneck in anti-glycan mAb production. Specifically, the use of microarrays enabled 50 μl of hybridoma supernatant to be screened rapidly and simultaneously against >60 potential epitope-bearing target molecules. A novel aspect of this work was the use of cluster analysis of array data to rapidly predict antibody specificities by comparison with previously defined mAbs. Subsequent detailed analyses of the specificities of the new mAbs LM13 and LM14 indicated they bound to pectic and AGP class of polymers respectively, as was predicted by the cluster analysis. These results indicate that if a relatively large set of probes with defined specificities are available to serve as references, this is an effective method for high-throughput initial mAb screening. One potential limitation of shotgun immunisation could be immuno-dominance, such that an immune response is elicited against a limited subset of the injected antigens. In this study two mAbs, LM13 and LM14 were selected with specificity for two different classes of molecule. However, of the seven mAbs selected for cluster analysis of binding profiles, six clustered with mAbs with specificity for AGPs and one with mAbs to pectic side chains, suggesting that AGPs were the immuno-dominant antigens in this case. It is therefore likely that such a multi-antigen approach towards cell wall polymers may be most effective to obtain a panel of mAbs with a range of specificities within a single class of polymer.
The arrays we constructed were based on the non-covalent attachment of glycans to nitrocellulose. This approach has the advantage that molecules can be immobilized directly without the need to create functional groups, and nitrocellulose has been previously shown to be a effective substrate for the immobilization of diverse glyans [35, 36]. Non-covalent attachment has the potential drawback that variations in the effectiveness of immobilization may result from differences in the structural properties of the arrayed molecules [8]. All of the arrayed samples used in this work were recognized to some extent by at least one of the mAbs used (as shown in Fig. 4), indicating that all were immobilized to some degree. Nevertheless, the spot signals obtained can only provide semi-quantitative information about mAb binding.
Two new probes, LM13 and LM14 were produced against cell wall polymers and both have distinctly different specificities to previously generated antibodies. Cluster analysis, ciELISA data immunoblotting and arabinanase sensitivity suggested that the epitope recognized by LM13 is an arabinan-containing structure that occurs as a side chain on pectic polymers. Pectic side chains typically consist of either galactan, arabinan or type I arabinogalactan chains in which arabinose usually occurs as a terminal sugar [18]. It was of note that LM13 binding to arrays was reduced to some extent by galactanase digestion. However, the galactanase used had very low arabinanase side activity (<0.03 U/ml specific activity with an arabinan substrate compared to 780 U/ml specific activity with a galactan substrate). It is possible therefore that galactanase digestion resulted in the indirect loss of the LM13 epitope by cleavage of galactan to which an arabinan-containing epitope is attached. The fact that LM14 bound to both pectin-derived MHR samples and to AGP-like material on a blot suggests that this mAb binds to an epitope of type II arabinogalactan that may occur on both pectins and AGPs. The novel patterns of recognition on A. thaliana and other plant materials indicates that these are useful new probes for the analysis of cell wall glycans polymers and complex cell wall architectures.

monoclonal antibodies
plant cell walls
hierarchical clustering
carbohydrate microarrays


The atypical iron-coordination geometry of cytochrome f remains unchanged upon binding to plastocyanin, as inferred by XAS

The transient complex between cytochrome f and plastocyanin from the cyanobacterium Nostoc sp. PCC 7119 has been analysed by X-ray Absorption Spectroscopy in solution, using both proteins in their oxidized and reduced states. Fe K-edge data mainly shows that the atypical metal coordination geometry of cytochrome f, in which the N-terminal amino acid acts as an axial ligand of the heme group, remains unaltered upon binding to its redox partner, plastocyanin. This fact suggests that cytochrome f provides a stable binding site for plastocyanin and minimizes the reorganization energy required in the transient complex formation, which could facilitate the electron transfer between the two redox partners.
Introduction
In oxygen-evolving photosynthetic organisms, the cytochrome b6–f complex couples proton translocation across the thylacoid membrane to the electron transport between photosystems I and II (Allen 2004). In this complex, cytochrome f (Cf) transfers electrons from the Rieske iron sulphur cluster to a soluble metalloprotein that acts as the immediate electron donor of P700 cofactor at Photosystem I.
The Cf subunit consists on a 28.2 kDa N-terminal soluble domain anchored to the membrane by C-terminal helix (Gray 1992). Its soluble domain is an atypical c-type cytochrome because both its β-sheet secondary structure and the unusual heme axial coordination (Martinez et al. 1994) with the N-terminal amino acid Tyr-1 acting as an axial ligand.
The most ubiquitous electron carrier between Cf and P700 is plastocyanin (Pc) (Sandman et al. 1983), which is a type I cupredoxin (Gough and Chotia 2004). Its structure consists on an anti-parallel β-barrel with a single copper atom (Coleman et al. 1978; Sykes 1985; Redinbo et al. 1994) coordinated by two nitrogen atoms and two sulphur ones from highly conserved residues (H39, C89, H92 and M97 in Nostoc cyanobacterium).
The mechanism of the electron transfer (ET) reaction between Pc and its physiological partners has been studied extensively (Hope 2000; Hervás et al. 2003; Díaz-Quintana et al. 2003) highlighting the role of electrostatic and hydrophobic interactions on binding. Also, the solution structures of several Pc–Cf complexes have been reported (Ubbink 2004; Díaz-Moreno et al. 2005a, Lange et al. 2005; Musiani et al. 2005). In all the cases, these structures are consistent with a single conformation ensemble in which the hydrophobic patch surrounding Tyr-1 in Cf docks the hydrophobic patch of Pc.
Despite the large amount of data concerning the nature of the interactions influencing the binding between the redox partners, little is known about the effects of complex formation on the metal cofactors, and how they can modulate the ET process. An analysis about whether the Fe coordination in Cf is altered upon binding to Pc would be interesting to address the above-mentioned effects, especially due to the fact that the N-terminal amino acid Tyr-1, which is involved in the Pc–Cf interface (Ubbink 2004; Díaz-Moreno et al. 2005a, Lange et al. 2005; Musiani et al. 2005), acts as axial ligand. To get a deep insight on these subjects we have studied the Fe K-edge of free and Pc-bound Cf by X-ray absorption spectroscopy (XAS) in solution.
Materials and methods
Protein samples
Nostoc sp. PCC 7119 Pc was purified from E. coli cells transformed with the pEAP-WT plasmid (Molina-Heredia et al. 1998). Production and purification of the soluble domain of Nostoc sp. PCC 7119 Cf were as previously described (Albarrán et al. 2005).
For the XAS experiments, the Pc and Cf samples were concentrated to the required volume by ultrafiltration methods, and exchanged into 10 mM sodium phosphate pH 6.0. After concentration, stock solutions of 10 mM Pc and 4.5 mM Cf were obtained. Protein concentration was determined by optical spectroscopy using an absorption coefficient of 4.5  mM−1 cm−1 at 598 nm for oxidized Pc (Molina-Heredia et al. 1998) and 31.5 mM−1 cm−1 at 556 nm for reduced Cf (Albarrán et al. 2005). Oxidized Cf (Cf III) was prepared by adding an equimolecular amount of potassium ferricyanide, followed by gel filtration in a Pharmacia Superdex G75 column to remove ferrocyanide. Reduced Cf (Cf II) was obtained upon addition of a tenfold excess of sodium ascorbate to a resulting Cf III sample. Oxidized Pc (PcII) was obtained by addition of either ferricyanide, as described for Cf. The oxidized complex (PcII–Cf III) was prepared by adding an aliquot of a 10 mM PcII onto a Cf III sample up to reach a final concentration of ca. 2 mM each. Under these conditions, the percentage of Pc bound to Cf was estimated to be ca. 87%, according to the binding constant calculated from NMR data (Díaz-Moreno et al., 2005a). The reduced complex (PcI–Cf II) was obtained upon addition of a tenfold excess of sodium ascorbate to a PcII–Cf III sample prepared as above.
XAS measurements
The X-ray absorption spectra were recorded at the European Synchrotron Radiation Facility (ESRF) in Grenoble (France), which was operating with a ring current of 200 mA and energy of 6 GeV. The Fe K-edge (7112 eV) was measured at beam line BM29 using a double crystal monochromator fitted with a pair of flat Si (311) crystals detuned to 50% fwhm of the maximum transmission, for the suppression of high energy harmonics.
All measurements were made at room temperature in fluorescence mode, using a 13-element Canberra solid-state germanium detector. The samples were contained in a PTFE cell equipped with two 12 μm Kapton foil windows.
Energy calibration was achieved by measuring an iron foil for the Fe edge. The spectra of the foil were measured simultaneously, with the data in transmission mode, to be used as an internal reference. Ionization chambers filled with the appropriate mixture of gases were used as detectors.
For all measurements, each data point was collected for 4 s, and several scans were averaged to achieve a good signal-to-noise ratio. In no case protein photoreduction or damage was observed, even in those samples in which the measurements were made over several hours. For the oxidized forms free and Pc-bound Cf, the edge remained at fix energy position from the first to the last scan, thus confirming the absence of protein photoreduction. The lack of radiation-induced changes in the samples was also confirmed by UV-vis measurements before and after XAS measurements.
XAS data analysis
The background subtraction required to obtain the extended X-ray absorption fine structure (EXAFS) functions χ(k) from the measured X-ray absorption spectra were performed using the AUTOBK code from the University of Washington. E0 was defined as the maximum of the first derivative of the absorption edge.
To analyze the EXAFS spectra, the theoretical phases and amplitudes were calculated using the FEFF 6.0 code (Rehr et al. 1992; Newville et al. 1995). The fit to the experimental data was performed using the FEFFIT program (version 2.54) (Newville et al. 1995).
Results and discussion
XANES region
Figure 1 (upper panel) shows the X-ray absorption near edge structure (XANES) spectra corresponding to free Cf in its reduced and oxidized states. In both spectra there is a pre-edge feature corresponding to the forbidden transition 1s−3d, which is sensitive to the electronic and geometric structure of the atom (Fe) site (Westre et al. 1997). In fact, the intensity of this transition agrees with the octahedral environment around the iron atom in Cf (Martinez et al. 1994).
Fig. 1XANES region of the Fe K-edge XAS spectra of free and Pc-bound Cf. Upper, experimental data for oxidized and reduced forms of free proteins, Cf III (continuous line) and Cf II (dashed line). Middle, data for the oxidized species, Cf III (continuous line) and PcII–Cf III complex (dashed line). Lower, data for the reduced species, Cf II (continuous line) and PcI–Cf II complex (dashed line). The presence of a pre-edge signal at 7113 eV is marked by arrows
On the other hand, the edge position, determined by the maximum in the first derivative of the absorption spectrum corresponding to free Cf II, appears ca. 1 eV below that of Cf III, as previously found for the reduced and oxidized states of two closely related proteins, as heart cytochrome c (Cheng et al. 1999) and Nostoc cytochrome c6 (Díaz-Moreno et al. 2006). In case of cytochrome c (Cheng et al. 1999), photoreduction of the oxidized species was observed only after the 16th spectrum of the series. In our case, however, there is no photoreduction as inferred from the fact that the edge position remains fixed for all the spectra along each series i.e. reduced or oxidized, free or bound. The reason not to observe any sample photo-reduction is due to the moderate brightness of the EXAFS spectrometer deliberately chosen for this investigation. In addition, the same shift has also been reported upon oxidation from Fe2+ to Fe3+ at pH 7 in the heme protein cytochrome c (Shulman et al. 1976).
Moreover, there are some differences in both XANES spectra, particularly in the first feature after the edge.
Figure 1 also shows the XANES regions of the absorption spectra of Cf bound to Pc in the oxidized and reduced state. The XANES region of the absorption spectra for PcII–Cf III and Cf III are identical (Fig. 1, middle panel), indicating that the electronic and geometrical configuration around the iron centre remains unchanged when bound to Pc. However, this is not the case for the reduced proteins (Fig. 1, lower panel). In fact, the spectrum of the PcI–Cf II complex exhibits some differences compared with that of free CfII. With regard to the pre-edge, a slight increase in the intensity of this feature is observed for CfII when bound to PcI, according to the decrease in electron density of the Fe atom. The shape of the first feature after the edge slightly changes too, thus suggesting certain modifications in the geometrical distribution of the ligands around the iron atom. It is worth noting that the spectrum of the reduced complex is very similar––except for the pre-edge signal and edge position––to that of the oxidized complex and free oxidized Cf, as shown in Fig. 3. This suggests that the iron geometry in reduced Cf is distorted upon association to Pc, approaching to that of the iron centre in the oxidized form, either free or bound to Pc.
EXAFS Region
The extracted EXAFS signals of Cf in the reduced and oxidized states are shown in Fig. 2 (left panels) as k-weighted spectra. The choice of a k-weighting instead of a k3-weighting is based on the following facts. First, we were interested in determining the local environment around the absorbing atom and, in particular, at the first coordination shell. This shell is formed by iron and light elements, such as nitrogen, and their backscattering amplitude drops quickly at increasing energy (k value). Therefore, they have a larger contribution when the transforms are made in k rather than in k3. Second, if a k3-weighting is used, the highest energy part of the spectrum is emphasized and, in this case, the EXAFS signal would be dominated by the spectral region where the signal-to-noise ratio is poorer.
Fig. 2EXAFS data (panels a and b) and their FT modules (panels c and d) at the Fe K-edge of Cf, either free or bound to Pc. The EXAFS spectra of free proteins (Cf II and CfIII) are shown in panela by continuous line for Cf II and dashed line for Cf III. In panel b, the EXAFS spectra of PcI–Cf II and PcII–Cf III complexes are represented by continuous and dashed lines, respectively. The corresponding FT modules of the EXAFS spectra for free Cf (Cf II and CfIII) are represented in panel c by closed and open circles, respectively. Panel d shows the FT modules for Pc-bound Cf forms: PcI–Cf II by closed circles and PcII–Cf III by open circles. The best fits of the FT data for both reduced (Cf II and PcI–Cf II) and oxidized states (Cf III and PcII–Cf III) are represented by continuous and dashed lines, respectively
We should note here that the XAS data were collected at room temperature in order to maintain consistency with previous room temperature NMR measurements made on the same systems (Díaz-Moreno et al. 2005a, b). Although a better signal to noise should be obtained at cryogenic temperatures, the transient complexes we study are not stable upon freezing, unless the partners are chemically cross-linked. Similar experimental conditions are required by EPR. Although EPR measurements on free and Pc-bound Cf would help to study the iron geometry, any comparison between results inferred from NMR or XAS, on one hand, and EPR, on the other, would be unreliable.
As can be seen, the EXAFS function is not a simple wave but a combination of different waves corresponding to the contributions from the neighbouring atoms at different distances around the iron centre. Indeed, the EXAFS signals of Cf do not change upon binding to Pc, thereby indicating that the local coordination environment around the iron centre remains unchanged.
The Fourier Transform (FT) of the corresponding spectra and the best fits, obtained with the parameters included in Table 1, are also shown in Fig. 2 (right panels). As a starting model, the values obtained from the X-ray diffraction (XRD) structure of Phormidium Cf (Carrel et al. 1999) were used. In all cases there is a main signal around 1.5 Å, which corresponds to the first coordination sphere formed by the four nitrogen atoms of the porphyrin ring (NA, NB, NC and ND), along with the nitrogen atoms of the two iron axial ligands Tyr-1 (N) and His-26 (Nε2). The intensities of the remaining signals are significantly lower than the first one because they correspond to contributions from lightweight elements lying at longer coordination distances. In order to fit the data, six simple scattering paths from the Fe atom to the six ligand N atoms were selected. Δr1 and Δr2 were used as two fitting parameters corresponding to changes in the distances of Fe to the two axial N atoms and to the four phorphyrin N atoms, respectively. The Debye–Waller parameter, σ2, related to system dynamic disorder was also fitted, with σ12 accounting for the bonds between the Fe atom and the two axial N atoms and σ22 accounting for the bonds between the Fe atom and the four porphyrin N atoms. The fifth adjustable parameter was the internal potential correction (ΔE0).
Table 1Best-fit structural parameters resulted from the EXAFS analysis for free and Pc-bound Cf, in both oxidized and reduced stateSystemLigandΔE0 (eV)r(M–L) (Å)σ2(M–L) (Å2)CfIIIN (NC)−7 ± 41.96 ± 0.020.002 ± 0.001N (NA & ND)1.97 ± 0.02N (NB)1.98 ± 0.02N (Nε2)2.44 ± 0.060.010 ± 0.010N (N)2.49 ± 0.06CfIIN (NC)−6 ± 21.97 ± 0.010.002 ± 0.001N (NA & ND)1.99 ± 0.01N (NB)2.00 ± 0.01N (Nε2)2.43 ± 0.020.004 ± 0.003N (N)2.48 ± 0.02PcII–CfIIIN (NC)−7 ± 31.97 ± 0.010.002 ± 0.001N (NA & ND)1.98 ± 0.01N (NB)1.99 ± 0.01N (Nε2)2.40 ± 0.030.011 ± 0.009N (N)2.45 ± 0.03PcI–CfIIN (NC)−6 ± 31.97 ± 0.020.003 ± 0.001N (NA & ND)1.98 ± 0.02N (NB)1.99 ± 0.02N (Nε2)2.40 ± 0.030.010 ± 0.009N (N)2.45 ± 0.03S02, factor of amplitude of reduction; r(M–L), metal–ligand distanceS02 = 1.0, Δk = 3.5–11 (1/Å), ΔR = 1.1–2.1 (Å), R factor (free Cf III) = 0.044, R factor (free Cf II) = 0.004, R factor (PcII-bound Cf III) = 0.004, R factor (PcI-bound Cf II) = 0.003
Although there are no significant changes in the metal–ligand distances, the Fe K-edge X-ray absorption data suggests a small distortion of the Fe2+ metal centre geometry when Cf II binds to PcI. The resulting Fe2+ geometry resembles that of Cf III, either free or bound to PcII (Fig. 3). Although the Fe atom of Cf shows a well-defined position at the heme group, which provides a rigid frame, the small distortion on the Fe2+ metal centre when Cf II binds to PcI can be explained by the smaller size of Fe3+ compared to Fe2+: The Fe3+ atom fits well into the heme ring whereas the Fe2+ atom is somewhat shifted out of the plane (Schnackenberg et al. 1999); however, the Fe2+ atom could be driven into the ring upon complex formation, thus yielding a structure of PcI-bound Fe2+ similar to that of Fe3+ in free and PcII-bound Cf III.
Fig. 3XANES region of the Fe K-edge XAS spectra of free CfIII and PcI–bound CfII. Experimental data for Cf III and PcI–Cf II are shown in continuous and dashed lines. The presence of a pre-edge signal at 7113 eV is marked by arrows
This tiny perturbation is consistent with the intensity changes at the pre-edge region of CfII in presence of PcI, because they correspond to transitions to the iron dz2 and dx2y2 orbitals, which are pointing towards the axial and equatorial ligands, respectively. As the reducing electron is not located in these orbitals, such subtle changes may have little influence on the redox potential. This situation is clearly different to that previously reported for cytochrome c6, which shows a perturbation on the pre-edge region corresponding to its oxidized species, instead of the reduced ones, upon binding its reaction partner, photosystem I (Díaz-Moreno et al. 2006). In such case the pre-edge corresponds to transfer to a half-occupied non-bonding d orbital, which is that receiving the reducing electron, so the observed changes can be correlated with changes in the redox potential of the heme protein. Regarding to this point, it is important to note that the heme plane is normal to the interaction surface of cytochrome c6, but parallel in the case of Cf.
Thus, our results indicate that the bond between iron and the N-terminus nitrogen is strong enough to prevent a distortion of the iron-coordination geometry upon binding of Cf to Pc, as well as to provide a stable binding site to the copper protein. Such a stiffness of the coordination sphere minimises the reorganisation energy concomitant to charge transfer, thus optimising the ET process.

cytochrome f
plastocyanin
transient complexes
x-ray absorption spectroscopy
electron transfer
metalloproteins